Medicine, Faculty of
Biochemistry and Molecular Biology, Department of
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UBCV
Boissy, Robert James
Doctor of Philosophy - PhD
University of British Columbia
Two important experimental techniques used to study eukaryotic, sequence-specific DNA-binding proteins are electrophoretic mobility-shift assays (EMSAs) and sequence-specific DNA-affinity chromatography. This thesis describes some new approaches to these frequently used methods, which were used to study a DNA-binding protein (ceHSBF) with an apparent specificity for a conserved DNA sequence element (the heat-shock element or HSE) that mediates the transcriptional regulation of heat-shock-inducible genes in the soil nematode Caenorlzabditis elegans. Although C. elegans has attracted extensive research interest, a lack of appropriate and reliable experimental methods has severely limited biochemical studies of sequence-specific DNA-binding proteins from this species. As a first step towards addressing this problem, I developed a simple and effective procedure for isolating C. elegans nuclei. I also developed a new class of EMSA electrophoresis buffers and established the running conditions appropriate for their use. The distinctive electrolyte in these buffers is monopotassium hydrogen glutamate (KHG1u), which provides both a physiological concentration of potassium ions (100- 150 mM) and a counterion species (glutamate) that is well-known for its ability to stabilize protein-DNA interactions. Two protein-DNA complexes that appeared to be specific for an HSE-bearing probe were detected using C elegans nuclear extracts and the KHG1u-based EMSA system. These complexes were believed to have been formed by two distinct types of putative ceHSBF activity, which were designated as ceHSBF 0and ceHSBF. The Rf(-.0. 1-0.2) of the protein-HSE complex formed by the putative ceHSBF 0activity agrees with the known Rf of the complex formed by most of the homotrimeric heat-shock factors from other species. The Rf (—0.4-0.5) of the protein-HSE complex formed by the putative ceHSBFm activity suggests that this protein is either a discrete proteolytic degradation product of ceHSBF 0,or that ceHSBFm is a novel HSE-specific DNAbinding protein. The putative ceHSBFm activity is readily detectable in the presence of moderate concentrations of KC1, nonionic detergents, or a 200,000-fold mass-excess (relative to the labeled HSE probe) of the nonspecific competitor DNA poly(dI-dC).poly(dI-dC), but cannot be detected if the nuclear extract has been exposed to proteinase K or a 50-fold excess of unlabeled HSE probe DNA. In the second part of this thesis I examined various parameters that influence the covalent immobilization of unmodified, double-stranded DNA on acyl hydrazide-derivatized chromatography resins. Commercially available, derivatized agarose or poly(methyl methacrylate) resins were examined. A reaction involving the transainination of transiently single-stranded deoxycytidylate residues by acyl hydrazide groups of the resin is proposed to explain the immobilization. Some noteworthy properties of this immobilization chemistry include satisfactory yields and efficiencies, with 1.0 mg of a 2,686 base-pair fragment of DNA immobilized per millilitre of resin, with 80% efficiency, under standard reaction conditions (6 hours, 30-37 °C, pH 4.20); greater safety and convenience compared to CNBr-activated agarose resins; and the suitability of immobilized DNA for use as an affinity chromatography ligand, or as a solid-phase substrate for various physical, chemical and enzymatic reactions performed at ambient or elevated temperatures.
Some new approaches to conventional methodsfor the study ofprotein-DNA interactions.byRobert James BoissyB.Sc., The University of British Columbia, 1984.A THESIS SUBMITTED IN PARTIAL FULFILMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIESDepartment of Biochemistry and Molecular Biology.We accept this thesis as conformingto the required standard:THE UNIVERSITY OF BRITISH COLUMBIANovember, 1994.© Robert James Boissy, 1994.In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives, it is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)__________________________Department of 8iot/srA.y ,4Ahf L1Lji9- ‘oYThe University of British ColumbiaVancouver, CanadaDate ‘4/di’DE-6 (2/88)11ABSTRACTTwo important experimental techniques used to study eukaryotic, sequence-specific DNA-bindingproteins are electrophoretic mobility-shift assays (EMSAs) and sequence-specific DNA-affinitychromatography. This thesis describes some new approaches to these frequently used methods, which were usedto study a DNA-binding protein (ceHSBF) with an apparent specificity for a conserved DNA sequence element(the heat-shock element or HSE) that mediates the transcriptional regulation of heat-shock-inducible genes inthe soil nematode Caenorlzabditis elegans.Although C. elegans has attracted extensive research interest, a lack of appropriate and reliableexperimental methods has severely limited biochemical studies of sequence-specific DNA-binding proteins fromthis species. As a first step towards addressing this problem, I developed a simple and effective procedure forisolating C. elegans nuclei. I also developed a new class of EMSA electrophoresis buffers and established therunning conditions appropriate for their use. The distinctive electrolyte in these buffers is monopotassiumhydrogen glutamate (KHG1u), which provides both a physiological concentration of potassium ions (100-150 mM) and a counterion species (glutamate) that is well-known for its ability to stabilize protein-DNAinteractions.Two protein-DNA complexes that appeared to be specific for an HSE-bearing probe were detectedusing C elegans nuclear extracts and the KHG1u-based EMSA system. These complexes were believed to havebeen formed by two distinct types of putative ceHSBF activity, which were designated as ceHSBF0 andceHSBF. The Rf(-.0. 1-0.2) of the protein-HSE complex formed by the putative ceHSBF0activity agrees withthe known Rf of the complex formed by most of the homotrimeric heat-shock factors from other species. The Rf(—0.4-0.5) of the protein-HSE complex formed by the putative ceHSBFm activity suggests that this protein iseither a discrete proteolytic degradation product of ceHSBF0,or that ceHSBFm is a novel HSE-specific DNA-binding protein. The putative ceHSBFm activity is readily detectable in the presence of moderate concentrationsof KC1, nonionic detergents, or a 200,000-fold mass-excess (relative to the labeled HSE probe) of theI”nonspecific competitor DNA poly(dI-dC).poly(dI-dC), but cannot be detected if the nuclear extract has beenexposed to proteinase K or a 50-fold excess of unlabeled HSE probe DNA.In the second part of this thesis I examined various parameters that influence the covalentimmobilization of unmodified, double-stranded DNA on acyl hydrazide-derivatized chromatography resins.Commercially available, derivatized agarose or poly(methyl methacrylate) resins were examined. A reactioninvolving the transainination of transiently single-stranded deoxycytidylate residues by acyl hydrazide groups ofthe resin is proposed to explain the immobilization. Some noteworthy properties of this immobilizationchemistry include satisfactory yields and efficiencies, with 1.0 mg of a 2,686 base-pair fragment of DNAimmobilized per millilitre of resin, with 80% efficiency, under standard reaction conditions (6 hours, 30-37 °C,pH 4.20); greater safety and convenience compared to CNBr-activated agarose resins; and the suitability ofimmobilized DNA for use as an affinity chromatography ligand, or as a solid-phase substrate for variousphysical, chemical and enzymatic reactions performed at ambient or elevated temperatures.ivTABLE OF CONTENTSABSTRACTTABLE OF CONTENTS ivLIST OF TABLESLIST OF FIGURES viiiABBREVIATIONS, SYMBOLS, AND CONVENTIONSACKNOWLEDGMENTS AllDEDICATIONPART 1 1CHAPTER 1.1. INTRODUCTION 2A. Eukaryotic, sequence-specific DNA-binding proteins 21. Research significance 22. Detection, purification, and characterization 33. Commonly observed structural features 54. The recognition of specific DNA sequences 7a. The physico-chemical forces involved in protein-DNA binding 7b. The principle of molecular complementarity 8c. The kinetics of cognate binding-site selection 11d. Cognate binding-site selection in vivo 12B. Regulation of the transcription of eukaryotic heat-shock genes 131. The heat-shock response 132. Heat-shock gene promoters 163. Heat-shock transcription factor (HSF) 19a. Interspecies and intraspecies diversity 19b. Quaternary structure 19c. General properties of the HSF monomer 20d. The trimerization domain 20e. The DNA-binding domain 21f. The transcriptional activation domains 22g. The regulatory “masking” region 234. The biochemical mobilization and demobilization of HSF 24a. Signal transduction 24b. The response of HSF to heat shock 26c. An autoregulatory role for heat-shock proteins9 31C. The nematode Caenorhabditis elegans 341. Salient attributes 342. Studies of DNA-binding proteins and chromatin structure 353. Isolation of genes that encode sequence-specific DNA-binding proteins 354. Heat-shock-inducible gene expression 36D. Summary of thesis objectives for Part 1 37CHAPTER 1.2. MATERIALS AND METHODS 39A. Materials used in this thesis (Parts 1 and 2) 39B. Routine experimental procedures used in this thesis (Parts 1 and 2) 401. Preparation of solutions 402. Liquid handling and plastic ware 43V3. Aseptic que.444. Mixing by rotation 445. Small-scale degassing of solutions or chromatography resins 446. Microcentrifugation 447. Miscellaneous procedures for handling DNA 45a. DNA precipitation and resuspension 45b.DNA assays 46C. Preparation of nuclear extracts from C. elegans eggs 461. Large-scale liquid culture of C. elegans 462. Large-scale preparation of C. elegans eggs and exposure to heat shock 473. Preparation of 0.35 M KCI nuclear extracts: The A-protocol 494. Preparation of 0.35 M KC1 nuclear extracts: The X-protocol and the AP-protocol 515. Quantitative analyses of A-protocol nuclei and nuclear extracts 52D. Electrophoretic mobility-shift assays (EMSA5) 531. Oligonucleotides used to form HSE probes 532. Fill-in end-labeling of the HSE probe 553. Preparation and PAGE analysis of EMSA binding reactions 56CHAPTER 1.3. RESULTS 58A. Preparation of nuclei from C. elegans eggs 58B. Electrophoretic mobility-shift assays (EMSAs) 621. The X-protocol results: Detection of a putative ceHSBF0activity 622. The AP-protocol results: Detection of both types of putative ceHSBF activity 673. The A-protocol results: Detection of a putative ceHSBFm activity 69CHAPTER 1.4. DISCUSSION 77A. The study of sequence-specific DNA-binding proteins from C. elegans 77B. The detection of two types of putative ceHSBF activity 781. Difficulties in identifying ceHSBF activity 782. The putative ceHSBF0activity 813. The putative ceHSBFm activity 834. The putative HSF activity in the hehninth Schistosoma mansoni 86C. The utility of KHG1u-based electrophoresis buffers in EMSAs 861. EMSA electrophoresis buffers presently in use 862. The rationale for the formulation of KHG1u-based EMSA electrophoresis buffers 87PART 2 91CHAPTER 2.1. INTRODUCTION 92A. Cytosine-specific transamination reactions 921. Reactivity of the pyrimidine ring in cytosine derivatives 922. The Bucherer reaction 923. The deamination and transamination reactions of cytosine derivatives 954. The transamination of cytosine derivatives by acyl hydrazides 97a. The chemistry of acyl hydrazides 97b. The reaction of acyl hydrazides with cytosine derivatives 995. Reactivity of cytosine moieties in polynucleotides 100B. Cyanate esters, acyl hydrazides, and the immobilization of nucleic acids on solid supports 1011. Strategies for the immobilization of DNA on chromatography resins 1012. The use of CNBr-activated agarose for the immobilization of unmodified DNA 1013. The use of acyl hydrazide-derivatized agarose for the immobilization of oxidized RNA.. 102C. Summary of thesis objectives for Part 2 103CHAPTER 2.2. MATERIALS AND METHODS 104A. Materials and routine experimental procedures 104B. The immobilization of DNA on acyl hydrazide-derivatized solid supports 1041. Preparation of DNA for use in immobilization assays 104via. Very large-scale preparation of pUC 19 plasmid DNA.104b. Restriction endonuclease digests of DNA 105c. Preparation of poly(dI-dC).poly(dI-dC) 1072. Handling of small quantities of chromatography resins 1073. The solubiity of TCBA salts at pH 4.2 1084. The standard DNA-immobilization reaction 1085. Storage of immobilized DNA and the assay of linkage stability 111C. Immobilized DNA as an affinity chromatography ligand 1111. Construction of an HSE-affmity resin 1112. Microscale affinity chromatography 112D. Immobilized DNA as a substrate for enzyme-catalyzed reactions 113CHAPTER 2.3. RESULTS 115A. The immobilization of DNA on acyl hydrazide-derivatized solid supports 1151. Development of the microscale DNA-immobilization protocol 1152. Formulation of the coupling buffer 116a. The hydrogen-ion buffer and its counterion 116b. Other components of the coupling buffer 1213. The nature of the deoxycytidylate-acyl hydrazide linkage 1264. The stability of the deoxycytidylate-acyl hydrazide linkage 131B. Immobilized DNA as an affinity chromatography liganci 132C. Immobilized DNA as a substrate for enzyme-catalyzed reactions 136CHAPTER 2.4. DISCUSSION 140A. The immobilization of DNA on acyl hydrazide-denvatized solid supports 1401. Effective-concentration and catalytic effects 1402. The roles played by individual components of the coupling buffer 141a. The role of the hydrogen-ion buffer 141b. The role of tricarballylic acid’s countenon 143c. The role of polyethylene glycol 1443. Important unanswered questions about the immobilization chemistry 145a. The distribution and average number of attachment points per DNA fragment. .. 145b. The deamination and depurination of DNA at pH 4.2 145c. The possibility of acyl hydrazone formation at apurinic sites in DNA 147B. Applications using DNA immobilized on acyl hydrazide-derivatized solid supports 1511. Some properties of acyl hydrazide-derivatized solid supports 151a. Versatility, convenience, and safety 151b. DNA-binding capacities of Affi-Prep HZ and Affi-Gel HZ 151c. “Uncapped” or unreacted acyl hydrazides 151d. The rate of loss of immobilized DNA 1522. Immobilized DNA as an affinity chromatography ligand 1533. Immobilized DNA as a substrate for enzyme-catalyzed reactions 155REFERENCES 157vi’LIST OF TABLESTable 1. Functionally important structural features of some sequence-specific DNA-binding proteins 7Table 2. Hydrogen-ion buffered solutions formulated for this work 41Table 3. KHGIu and the binding of the Xenopus estrogen receptor to the estrogen response element (ERE) 89Table 4. Tautomeric equilibria of cytosine derivatives in aqueous solution 96Table 5. Influence of weakly acidic buffers on the covalent immobilization of DNA on Affi-Prep HZ 117Table 6. Influence of tricarballylic acid’s counterion on the immobilization of DNA on Affi-Prep HZ 118Table 7. Structure and relative efficiency of buffers used to immobilize DNA on Affi-Prep HZ 119Table 8. Influence of supplementary KC1 on the immobilization of DNA on Affi-Prep HZ 121Table 9. Influence of PEG 8000 on the immobilization of DNA on Affi-Prep HZ 121Table 10. Influence of critical cations on the immobilization of DNA on Affi-Prep HZ and Affi-Gel HZ 123Table 11. Structure, relative efficiency, and inhibitory potential of piperazine and cyclam 124Table 12. Influence of caudemer sequence on the immobilization of DNA on Affi-Prep HZ 127Table 13. Preparation of DNA-affinity chromatography resins 130Table 14. Loss of immobilized DNA after ten months 131vi”LIST OF FIGURESFigure 1. Geometric isomorphism of deoxyribonucleotide pairs in double-stranded B-DNA 10Figure 2. The A-protocol for the preparation of nuclei and nuclear extracts from C. elegans eggs 48Figure 3. DNA sequences of the oligonucleotides used to fonn HSE probes 54Figure 4. Overall purity of C. elegans nuclei isolated using the A-protocol 60Figure 5. Morphology of C elegans nuclei isolated using the A-protocol 61Figure 6. The “h2” Apa I fragment bearing two copies of the obsolete HSE consensus sequence 62Figure 7. EMSA detection of a putative ceHSBF0activity 63Figure 8. The influence of competitor nucleic acids and heparin on the putative ceHSBF0activity 65Figure 9. Tolerance of noniomc detergents by the putative ceHSBF0activity 67Figure 10. Enhanced detection of a putative ceHSBF activity 68Figure 11. The influence of KC1 concentration on the two types of putative ceHSBF activity 69Figure 12. The influence of electrophoresis buffer pH on the detection of the putative ceHSBFm activity 71Figure 13. The apparent sequence-specificity of the putative ceHSBFm activity 71Figure 14. Order-of-addition analysis of ceHSBFm sequence-specificity 72Figure 15. The influence of proteinase K and nonionic detergents on the putative ceHSBFm activity 74Figure 16. The Bucherer reaction 93Figure 17. The AE mechanism for the deamination and transamination of cytosine derivatives 94Figure 18. Structural features of acetic acid hydrazide (acethydrazide) 98Figure 19. The reaction of Girard’s reagent P with cytosine derivatives 99Figure 20. Protocol for microscale DNA-immobilization reactions 110Figure 21. Microscopy of Affi-Gel HZ beads containing immobilized DNA 125Figure 22. Bisulfite-catalyzed immobilization of DNA on polyacrylhydrazido agarose 128Figure 23. EMSA and SDS-PAGE analyses of HSE-DNA-affinity column fractions 133Figure 24. Restriction endonuclease digestion products obtained using immobilized DNA 137Figure 25. A hypothetical mechanism for DNA immobilization via acyl hydrazone formation 148ixABBREVIATIONS, SYMBOLS, AND CONVENTIONSA Adenosine. In this thesis A denotes deoxyadenylate residues in DNA.Absorbance at wavelength (where is given in nrn).A Angstrom(1OA=lnm).ABS Absorption.AcOH Acetic acid.ACS American Chemical Society.ADCC Antibody-dependent cellular cytotoxicity.AE Addition-elimination (reaction mechanism).AP Apunnic.ATP Adenosine triphosphate.BIS-TRIS bis-[2-Hydroxyethyljimino-tris[hydroxymethyl]-methane.bp Base pairs.BSA Bovine serum albumin.C Cytidine. In this thesis C denotes deoxycytidylate residues in DNA (or cysteine).%CBjs Degree of crosslinking with N, N’-methylene-bisacrylamide (% of total monomer weight).C-terminal Carboxy terminal (end of a peptide or protein).Cat no. Catalogue number.CB Coupling buffer (used for DNA-immobilization reactions; composition varies).ceHSBF Heat-shock element-specific DNA-binding protein from C. elegans.ceHSBFrn ceHSBF whose protein-HSE complex migrates midway between the origin and the probe.ceHSBF0 ceHSBF whose protein-HSE complex barely migrates past the origin.Ci Curie (The SI unit is Bq, or becquerel; 1 Ci = 37 GBq and 1 Bq =60 dpm).CNT Control.:: (2 colons) Indicates plasmid-bearing status (bacterial strain::plasmid).cyclam 1, 4, 8, 11-Tetraazacyclotetradecane.d Deoxy.Da Daltons.DABCO 1,4-Diazabicyclo[2.2.2] octane.DAPI 4,6-Diamidino-2-phenylindole.dia. Diameter.DIC Differential interference contrast (microscopy).DNA Deoxyribonucleic acid.DNase Deoxyribonuclease.dNTP Deoxynucleoside triphosphate.(dot) Indicates annealed polynucleotide strands (see note (ii) below).DTPA Diethylenetriamine-pentaacetic acid.DTr Dithiothreitol.E-64 L-trans-Epoxysuccinyl-leucylamido(4-guanidino)butane.EC Embryonal carcinoma.EDTA Ethylenediamine-tetraacetic acid.EGTA Ethyleneglycol-bis(3-aminoethyl ether) N,N,N’,N’-tetraacetic acid.EMSA Electrophoretic mobility-shift assay.EPPS N-[2-Hydroxyethyl]piperazine-N’-[3-propanesulfonic acid].(epsilon) Molar extinction coefficient at wavelength ? (where is given in nm).Et Ethyl.EtdBr Ethidium bromide (2,7-diamino-10-ethyl-9-phenylphenanthridiniurn bromide).EtOH Ethanol.g Grams.g Acceleration of gravity.G Guanosine. In this thesis G denotes deoxyguanylate residues in DNA.xGABA 4-Aminobutanoic acid.h Hours.h2, h32 DNA fragments bearing 2 or 32 copies of the obsolete HSE consensus sequence.HEPES N-[2-Hydroxyethyl]piperazine-N’-[2-ethanesulfonic acid].HIV Human immunodeficiency virus.HS Heat-shock(ed).HSBF (Eukaryotic) heat-shock element-specific DNA-binding protein.HSE (Eukaryotic) heat-shock (gene promoter) element.HSF (HSTF) (Eukaryotic) heat-shock (gene) factor (or transcription factor).HSP Heat-shock protein.- (hyphen) Indicates covalent linkage of nucleotides, e.g., dI-dC (see note (ii) below).I Inosine.I Ionic strength.KHG1u Monopotassium salt of L-glutamic acid.? (lambda) Wavelength.M Molar concentration.Mr Relative molecular mass.Me Methyl.MES 2-[N-Morpholino]ethanesulfonic acid.MEL Murine erythroleukemia.mm Minutes.n Number of determinations.N Any nucleotide. In this thesis, either A, C, G, or T.N-terminal Amino terminal (end of a peptide or protein).NMR Nuclear magnetic resonance.NRC No-resin control.usc Nonspecific competitor DNA.OAc Acetate.OG n-Octyl 3-D-glucopyranoside.OH Hydroxyl.p Phosphate.PAGE Polyacrylamide gel electrophoresis.pDNA Plasmid DNA.pdN5 5’-Phosphorylated, random-sequence deoxyribopentanucleotides.pdN6 5’-Phosphorylated, random-sequence deoxyribohexanucleotides.PEG 8000 Polyethylene glycol (average Mr = 8000).PMSF Phenylmethanesulfonyl fluoride.pol II RNA polymerase II (B).N’ (psi) Polymer-and-salt-induced transition of DNA.pUC A series of plasmid cloning vectors.Rf Retardation factor.RNA Ribonucleic acid.rpm Revolutions per minute.s Seconds.SB Stop buffer (used to stop DNA-immobilization reactions; see below).SDS Sodium dodecyl sulfate.a (sigma) Bacterial transcription initiation factor (isoforms are designated by superscripts).STE NaC1 (100mM), TRIS-HC1 (10 mM),Na2EDTA (1 mM), pH 7.4 or 8.0.t Time.T Thymidine. In this thesis T denotes deoxythymidylate residues in DNA.The amount of total acrylamides as % w/v in a PAGE gel.xiTCBA Tricarbaflylic acid.Td Oligonucleotide thermal dissociation temperature.TE TRIS-HC1 (10 mM), Na2EDTA (1 mM), pH 7.4 or pH 8.0.TEMED N,N,N’,N’-Tetramethylethylenediamine.TEP TE containing 2% wlv PEG 8000.(tilde) Approximately.Tm Helix-coil transition temperature for the thermal denaturation of double-stranded DNA.TRIS tris[Hydroxymethyl]aminomethane.tRNA Transfer ribonucleic acid.Tx-100 Triton X-100 (a nonionic detergent).U Units of enzyme activity.UV Ultraviolet.v Volume.v:v Volume to volume (e.g., 5:1 v:v of buffer A.02:B).v/v Volume in volume (e.g., 10% v/v “x” = 10 mL of x per 100 mL of solution).vol Volumes.w Weight.w/v Weight (mass) in volume (e.g., 10% wlv “y” = 10 g of y per 100 mL of solution).The following were not listed:(1) SI units and prefixes.(2) Abbreviations and symbols used for systematic chemical nomenclature.(3) Abbreviated names for some of the buffers formulated for this work.See Table 2 in the Materials And Methods Section of Part 1 of this thesis.(4) Abbreviations for genetic loci (e.g., dnaK, etc.) or tissue-culture cell lines (e.g., HeLa, etc.).(5) Names assigned to transcription factors or other sequence-specific DNA-binding proteins(e.g., TFIID, Spl, etc.).(6) Restriction endonuclease names (e.g., Kpn I, Xba I, etc.).(7) Amino acid abbreviations. These are listed in (1985) J Biol. Chem. 260, 14-42.(8) Company names, brand names, or product model designations (e.g., HB-4, RC5B, IEC, etc.).Please also note that:(i) The ring numbering system used for cytosine is that recommended by the IUPAC and the IUB.This system is described in (1986) J. Biol. Chem. 261, 13-17.(ii) Double-stranded oligonucleotides are denoted using the JUPAC-JUB conventions described in (1970)J Biol. Chem. 245, 5171-5176.xl’ACKNOWLEDGMENTSThere were many people in the Biochemistry Department who allowed me to use facilities andequipment for which they were responsible, and who helped me order supplies. I offer them my sincere thanks.I also wish to express my gratitude to Oligos Etc. Inc., and in particular to David Pritchard and New EnglandBiolabs Ltd., for selling me the reagents required for the completion of my experimental work. NEB providedme with reagents at greatly reduced prices, and allowed me to defer payment until after I finished myexperimental work.I am also thankful for the help I received from the departmental office staff, the staff of variousbranches of the UBC Library, the staff of the UBC Biomedical Communications Department, and the staff of thePersonal Computing Support Center of the University Computing Services (UCS). This thesis was producedusing a very helpful set of UBC thesis formatting styles for use with the Microsoft Word word processingprogram. This formatting package was prepared by Eldon Wong of the UCS.My mother has always been a source of great encouragement and love, and I am very grateful for themany ways she helped me. My father, who died in 1969, lives on in my heart, and I will never forget how hardhe worked to provide for our family, and how highly he valued education. There are many other relatives,friends and colleagues to whom I am personally indebted, and I apologize for not being able to mention them all.I also apologize in advance for the inevitable errors and oversights in this document, especially the inadequatecitation of those scientists whose work is germane to this investigation.British Columbian and other Canadian taxpayers provided limited funding for my post-graduateeducation. I am grateful for this support. I am also grateful for the generous fmancial support I received frommy wife, Lai Chui Kwan, and from my father-rn-law, Mr. Lai Wai Hum of Hong Kong. I would not have beenable to complete my studies without their assistance and encouragement.xl”DEDICATIONLai Chui Kwan.Thoughtfulness colours her visionKindness animates her heartAnd the winter snowFears the warmth ofher smile.Trust engenders her commitmentFaith sustains her concernAnd the spring blossomsEnvy the beauty of her soul.Virtue enhances her wisdomLaughter graces her speechAnd the summer sunMarvels at the radiance of her spirit.Love inspires her compassionHope reaffirms her goodwillAnd the autumn leavesFall like my weary heartfor her own.- R.J.B.1PART 12CHAPTER 1.1. INTRODUCTIONA. EUKARYOTIC, SEQUENCE-SPECIFIC DNA-BINDING PROTEINS.1. Research significance.The development of DNA cloning and sequencing technologies has led to the discovery of a variety ofconserved DNA regulatory sequences that function during the replication, transcription, and rearrangement ofeukaryotic genomes (Faisst & Meyer, 1992; Held & Heinz, 1992; Leung & Maizels, 1992). A large number ofthese regulatory sequences have been shown to act as cognate (functionally related) binding sites for sequence-specific DNA-binding proteins (Faisst & Meyer, 1992; Freemont et al., 1991; Steitz, 1990). These sequence-specific protein-DNA interactions regulate various programs of gene expression necessary for development,survival, and reproduction (Bell & Stifiman, 1992: Pankratz et at., 1990; Rosner et at., 1990).Conversely, it is increasingly apparent that predisposition to developmental abnormalities, neoplastictransformation, and other serious diseases may result if particularly important members of this class of proteinsdo not function properly (Gessler et at., 1990; Goddard et at., 1991; Mellentin et at., 1989; Xia et at., 1991).The expression products of many oncogenes and anti-oncogenes (tumour-suppressor genes, i.e., genes whoselack of expression predisposes an individual to cancer) are believed to act as (or in conjunction with) sequence-specific DNA-binding proteins (Dressler & Douglass, 1992; Green, 1989; Haupt et at., 1991; Kim et al., 1992;Lee et al., 1987; Rauscher et al., 1990; Whyte et at., 1988). Programs of gene expression may also becompromised when invasive genetic information (propagated by viruses or mobile elements) directs theexpression of sequence-specific DNA-binding proteins that inactivate, exploit, or supplant the activities of theirhost-encoded counterparts. (Kekul6 et at., 1990; Lillie & Green, 1989; Seto et at., 1990).The most extensively studied eukaryotic, sequence-specific DNA-binding proteins are those involvedin the regulation of transcription, and in particular the transcription by RNA polymerase 11(B) of protein-codinggenes (Johnson & McKnight, 1989; Latchman, 1990; Mitchell & Tjian, 1989; Pabo & Sauer, 1992). Thedetection, purification, and analysis of these “transcription factors”, and the identification of the genes encodingthem, are exciting experimental challenges that have attracted a tremendous amount of interest (Faisst & Meyer,31992; Ghosh, 1991). However, there are some fundamental difficulties involved in pursuing these experiments.It is beyond the scope of this discussion to review the many technical approaches that have been developed toovercome these difficulties. Nevertheless, some knowledge of the techniques that are most commonly used isessential for a critical evaluation of the insights that these experiments have yielded.2. Detection, purification, and characterization.One problem associated with the study of sequence-specific DNA-binding proteins is that they arefrequently present in low abundance, especially if the protein of interest is inducible, developmental stagespecific, or cell type specific (Kadonaga, 1991). Another fundamental experimental difficulty is the propensityof sequence-specific DNA-binding proteins to bind DNA nonspecifically (Lohrnan, 1986; Record et al., 1991;von Hippel et al., 1984). Together these two problems present very serious obstacles, as the concentration andnumber of other nonspecific DNA-binding activities present in a whole-cell or nuclear extract (the latter extracttypically obtained using about 0.4 M NaC1) are considerable (Kadonaga, 1991). Other problems that may beencountered during the purification and analysis of sequence-specific DNA-binding proteins include (i) theleakage of nuclear proteins during subcellular fractionation; (ii) difficulties associated with assaying theprotein’s activity (or activities); (iii) protein or nucleoprotein aggregation; (iv) inthnsic protein lability;(v) nonspecific adsorption to surfaces; (vi) sensitivity to proteases; (vii) the loss of posttranslationalmodifications; and (viii) the existence of multiple protein isoforms (Kadonaga, 1991; Paine et al., 1983).Despite these difficulties, whole-cell or nuclear extracts are frequently used for the detection ofsequence-specific DNA-binding proteins, and for the mapping of protein-DNA contact points at nucleotideresolution. The most commonly used method for the detection of sequence-specific DNA-binding proteins is anelectrophoretic mobility-shift assay (EMS A) (Cann, 1993; Carey, 1991; Fried & Crothers, 1981; Garner &Revzin, 1981; Lane et al., 1992; Stone et al., 1991). In this technique a radiolabeled DNA probe containing thebinding site of interest is incubated with a whole-cell or nuclear extract (or purified protein), and the (typicallyless mobile) protein-DNA complex is separated from the free probe by nondenaturing, polyacrylamide gelelectrophoresis (PAGE). A crucial test of sequence specificity is the ability of a moderate excess of unlabeled,sequence-specific probe DNA—typically a 50- or 100-fold excess—to specifically compete off the4DNA-binding activity and abolish the autoradiograph signal of the protein-DNA complex. Conversely, a muchlarger excess of a nonspecific competitor DNA—typically a several thousandfold excess of a synthetic,alternating copolymer such as poly(dI-dC)•poly(dI-dC)—is shown to have little or no effect (Carthew et at.,1985; Singh et at., 1986).The purification of eukaryotic DNA-binding proteins of known sequence-specificity can be aparticularly daunting challenge. In the usual case (i.e., purification of a low-abundance regulatory protein) thenumber of purification steps should be kept to a minimum, and conventional-scale methods are frequentlyinappropriate. The penultimate purification step in most experiments is sequence-specific DNA-affinitychromatography (Kadonaga & Tjian, 1986; Rosenfeld & Kelly, 1986). This technique requires an affinitycolumn containing DNA fragments bearing the (known) cognate binding site of the protein of interest. In mostcases duplex oligonucleotides are ligated to form multiple repeats and then immobilized on a beaded agarosesupport using cyanogen bromide activation (Arndt-Jovin et at., 1975; Kadonaga, 1991; Kadonaga & Tjian,1986).So problematic is the purification of eukaryotic DNA-binding proteins of known sequence-specificitythat many researchers have resorted to isolating the gene(s) encoding a DNA-binding activity from a phageexpression library directly, i.e., based on the ability of the expressed gene product to bind to a radiolabeled DNAprobe containing the cognate binding site in question (Singh et at., 1988; Vinson et at., 1988). However, even ifsuch attempts are successful, and the cloned gene can subsequently be expressed in a bacterial or eukaryoticexpression system, the nature and functional significance of any posttranslational modifications remainunresolved. Such modifications are commonly observed, especially with inducible, sequence-specificDNA-binding proteins, or sequence-specific DNA-binding proteins with inducible functions, e.g., transcriptionalactivation (Jackson & Tjian, 1988; Sorger & Petham, 1988). In such cases an understanding of the nature andsignificance of posttranslational modifications may provide crucial insights into various mechanisms of signaltransduction (Berk, 1989).Examining the ability of a sequence-specific DNA-binding protein to protect a DNA sequence fromcleavage by deoxyribonuclease I (DNase I) is a commonly used method for the low-resolution mapping of5protein-DNA interactions (Galas & Schmitz, 1978). Chemical interference “footprinting” of sequence-specificDNA-binding proteins is a related technique for the mapping of protein-DNA contact points at nucleotideresolution (Siebenlist & Gilbert, 1980). In the latter experiment, a DNA probe is radiolabeled on one strand andthen modified with a chemical reagent under conditions where the average number of modifications per DNAmolecule is much less than one. An EMSA is performed, and then DNA samples from the protein-DNAcomplex and free probe bands are isolated. These samples are then treated with pipendine under conditionswhere the DNA strand is cleaved at those positions where the DNA had been chemically modified. Equalamounts of each DNA sample are then analyzed by denaturing PAGE. The signal intensity arising as a result ofcleavage at a given nucleotide residue should be smaller in the protein-DNA complex DNA lane (compared tothe intensity in the free probe lane) if that residue participates in sequence-specific recognition by theDNA-binding protein under study.X-ray crystallography and nuclear magnetic resonance (NMR) have also been used to study sequence-specific DNA-binding proteins, and in some cases the protein under study has been examined in a complex withits cognate DNA-binding site (Marmorstein et al., 1992; McClarin et a!., 1986; Párraga & Klevit, 1991; Russu,1991). These techniques can provide very high-resolution structural information, and X-ray crystallography hasproven extremely valuable for the study of prokaryotic sequence-specific DNA-binding proteins. Furthermore,it should be noted that many of the insights obtained from the study of prokaryotic systems have proven veryrelevant to our understanding of eukaryotic, sequence-specific DNA-binding proteins. However, both X-raycrystallography and NMR studies require large amounts of purified protein. In most cases the molecular cloningand subsequent overexpression of the gene encoding a sequence-specific DNA-binding protein are required toobtain enough material for these high-resolution structural studies.3. Commonly observed structural features.Genetic, biochemical, and physical studies of various sequence-specific DNA-binding proteins haveidentified several commonly observed structural features. These features include the ability to formsupraquaternary structures in vivo, e.g., the multiprotein ensembles involved in the regulation of transcriptioninitiation. At the level of primary structure, these features also include a variety of posttranslational6modifications. However, posttranslational modifications and multiprotein complex formation are frequentlyunrelated to sequence-specific DNA-binding per Se. The following discussion will focus on those structuralfeatures that are involved in sequence-specific DNA-binding, and not on those structural features involved incatalysis or its regulation.Many sequence-specific DNA-binding proteins bind to their cognate DNA-binding sites assymmetrically oriented homodimers (Pabo & Sauer, 1984; Schleif, 1988). The prevalence of this form ofquaternary structure is understandable given that DNA regulatory sequences are frequently palindromest withtwofold symmetry. The binding of dimer subunits is frequently cooperative, and this cooperativity effectivelyreduces the concentration of monomer required to achieve occupancy of both halves of the binding site(Kowalczykowski et a!., 1986a, 1986b). The formation of symmetrically oriented dimers may also be requiredfor certain regulatory sequences to act in a bidirectional manner. Heterodimerization also occurs, andheterodimer formation may provide a source of regulatory flexibility for some genetic loci (Lamb & McKnight,1991; O’Shea eta!., 1989; Shemshedmi et a!., 1991).The tertiary structure of sequence-specific DNA-binding proteins is characterized by modular domains(Frankel & Kim, 1991). These domains are usually involved in either DNA-binding, dimerization, ligandbinding, catalysis, or regulation (see Table 1). The construction and expression of chimeric genes hasdemonstrated the interspecies modularity of these domains (Agre et a!., 1989; Keegan et a!., 1986). Thisdomain modularity makes it important to distinguish the DNA-binding activity of a sequence-specificDNA-binding protein from some other associated activity. In some cases more that one protein can bindspecifically to a given binding site (e.g., Henthorn et a!., 1990). In other cases one protein is able to bindspecifically to more than one cognate binding site (e.g., restriction endonucleases; see Roberts & Macelis, 1993).In yet other cases one protein is able to bind to copies of the same binding site present at different loci and thenmediate different effects, presumably due to site occlusion or locus-specific protein-protein interactions (e.g.,Foulkes & Sassone-Corsi, 1992; Malcela eta!., 1992; Miner & Yamamoto, 1991; Westwood eta!., 1991).t DNA palindromes are double-stranded inverted repeats (e.g., GAATTC). The two antiparallel,complementary sequences that make up a DNA palindrome are identical.7Table 1. Functionally important structural features of some sequence-specific DNA-binding proteins.Structural feature Comments ExampleAmino tenninus Extended, flexible N-terminus penetrates the major ? repressorgroove. (Pabo et al., 1982).[3 Ribbon Antiparallel 13-strands that can bind in two orientations MetJ repressorto either the major or minor groove. (Somers & Phillips, 1992).Dinucleotide Major groove recognized by the amino ends of a-helices EcoR I(Rossman) fold that are part of a parallel [3-sheet. (McClarin et al., 1986).ETS domain DNA-binding domain consisting of an 85 amino acid ETS-1region of uncharacterized 2° structure. (Nye et al., 1992).Helix-loop-helix Dimerization interface with two short cx-helices Maxseparated by a loop region. (Blackwood & Eisenman, 1991).Helix-turn-helix DNA-binding domain consisting of two cx-helices ? repressorseparated by a 13-turn. One helix lies in the major (Harrison & Aggarwal, 1990).groove, and the other lies across this recognition’ helix.Homeodomain DNA-binding domain that is similar to the helix-turn- Engrailedhelix. (see Treisman et al., 1992).(HX) repeats Postulated to function as a pH-sensitive “histicline-metal Daughterlesszipper”. (see Janknecht et al., 1991).“Leucine zipper” Amphipathic a-helix with leucine heptad repeats. Two C/EBPdomain leucine zipper domains mediate dimerization by forming (Landschulz et al., 1988).a parallel coiled-coil structure.POU domain 80 amino acid region of uncharacterized 2° structure Pit-i, Oct-i, Unc-86 (i.e., POU)adjacent to a POU homeoclomain. (see Herr et al., 1988).RxxRxRxxC Part of a DNA-binding domain of uncharacterized 2° ReI/xB familysequence structure. Implicated in the redox regulation of v-Rel. (Kumar et al., 1992).SPXX sequence Found in nonspecific and sequence-specific DNA- Krüppelbinding proteins. Forms a 13-turn I stabilized by two (see Suzuki, 1989).hydrogen bonds.“Zinc fingers” DNA-binding domain. Typically 2 to 37 repeats of a TFIIIA(Cys2Hi) structure containing a two-stranded, antiparallel 13-sheet (Miller et al., 1985).and an a-helix. A central, firmly bound zinc atomforms a cys2hi tetrahedral coordination complex.Zinc-binding Typically one or two repeats of a zinc-finger-like Gal 4domain (Cys) structure with four to six cysteines per domain. (Marmorstein et al., 1992).4. The recognition of specific DNA sequences.a. The physico-chemicalforces involved in protein-DNA binding.Four types of forces influence the binding of proteins to double-stranded DNA (Goldman & Ollis,1990). Electrostatic forces between negatively charged DNA phosphates and positively charged amino acid side8chains (lysine, arginine, and to a lesser extent histidine residues) typically contribute 40 kJ moi1 per salt-bridgeto the stability of a protein-DNA complex. These forces are affected by the solvent, relatively long range(decreasing in magnitude in proportion to the square of the distance between the charges), and orientationindependent. Hydrogen bonds are relatively short-range forces (decreasing in magnitude in proportion to thecube of the distance between the participants), and are strongly affected by the degree of colinearity of the axisof the lone-pair orbital and the hydrogen-heteroatom bond. Properly formed hydrogen bonds contribute little tothe free-energy of binding (i.e., they simply replace solvent hydrogen bonds). However, the failure of solvent-inaccessible residues in a protein-DNA complex to adequately replace solvent hydrogen bonds carries asignificant penalty, and it is for this reason that hydrogen bonding is an important determinant of specificity(von Hippel & Berg, 1986). Two other forces that stabilize protein-DNA interactions are the hydrophobic effect(entropically favourable exclusion of ordered water from nonpolar surfaces) and van der Waals interactions(extremely short-range, transient dipole-induced dipole interactions that decrease in magnitude in proportion tothe sixth power of the distance between the participants) (Goldman & Ollis, 1990; Ha et al., 1989).b. The principle of molecular complementarity.The principle of molecular complementanty states that the specificity of protein-DNA binding isproportional to the ability of the protein and its cognate DNA-binding site to form a complex that maximizes thefree-energy gain attributable to the stabilizing forces described above (Harrison & Aggarwal, 1990). Hydrogenbonding between amino acid side chains and the edges of the heterocycic bases in the major groove of double-stranded B-DNA is a very important determinant of molecular complementarity. In this respect the two mostimportant properties of the major groove of B-DNA are its size and its ability to present a profile of atomscapable of hydrogen bonding and van der Waals interactions that is specific for each base pair (see Fig. 1). Thecorresponding profiles in the minor groove of B-DNA are less informative, and it is difficult to distinguish Tfrom C, CG from GC, or AT from TA due to geometric isomorphism (Seeman et al., 1976). The minor grooveof B-DNA is also too narrow (ranging from 5.7 A in random-sequence DNA to 3.5 A in AT-rich DNA) toaccommodate an a-helix, and it is too deep to allow amino acid side chains to simply extend from the protein inthe absence of any local secondary structure and form hydrogen bonds with the heterocyclic bases (Steitz, 1990).9N14.I t’i ‘-‘1’I II II II II II II II II II I56°C-G T-AOfHfrHNN-H...7N9 N9cJ1__o_.._ I‘4Ci. c :‘ciI I I I I I I II I I I I I I II I H I I I II I I I I I I II I I II I I II II I I IZLtH1 I II I I II I I II I I II II I I II I I I I I I II I I I I I I,)56° 10.7 A 54°çSJ 10.4 A 56°çI.,,JI II II II II II II II II II IHHHH0 C....HN-H — N?SN3’ 0 :c1I I I II I I II I I II I I II I I I10.4A10.7AG-C A-T10Figure 1. Geometric isomorphism of deoxyribonucleotide pairs in double-stranded B-DNA.Arrows indicate hydrogen bonding, with arrowheads directed at hydrogen-bond acceptors. The smallest arrowsrepresent Watson-Crick hydrogen bonding. The narrow and WIDE larger-size arrows represent potentialhydrogen bonding in the minor and MAJOR grooves, respectively. The thymine methyl group is representedby a large circle. The (Ci’ - Ci’ - Nx } angles and Ci’to Ci’ distances are approximations. Adapted fromSaenger (1983), Seeman et al. (1976), and Steitz (1990).The major groove of B-DNA is about 12 A wide and 6 to 8 A deep (Steitz, 1990). An a-helix is about12 A in diameter (including projecting side chains), and this form of secondary structure is commonly used byproteins to bind to DNA (Freemont et al., 1991; Pabo & Sauer, 1984; Schleif, 1988; Steitz, 1990). It is usuallythe N-terminus of the a-helix that extends into the major groove. The amino acid side chains all point towardsthis end of an a-helix, which is also the positive end of the a-helix dipole. This allows one face of the straight,cylindrical a-helix to present a series of amino acid side chains that can contact a maximum of four to sixdeoxyribonucleotide pairs in the curving spiral of the major groove. Although a-helices are clearly an importantform of secondary structure used by sequence-specific DNA-binding proteins to bind to DNA, other forms ofsecondary structure have also been identified that serve this purpose (see Table 1).There is no molecular code for the recognition of individual deoxyribonucleotide pairs by amino acidside chains (Pabo & Sauer, 1984; Steitz, 1990). A given deoxyribonucleotide pair can be recognized by morethan one amino acid side chain, and a given amino acid side chain can recognize more than onedeoxyribonucleotide pair. However, there are some preferences, and these typically involve bidentate hydrogenbonding, e.g., between arginine and guanine, glutamine and adenine, and in some cases between arginine orglutamate and two consecutive adenine moieties (Pabo & Sauer, 1984; Seeman et al., 1976; Steitz, 1990).Bidentate hydrogen bonding involving a given amino acid side chain, a deoxyribonucleotide pair, and anotheramino acid side chain is also observed (Steitz, 1990). In some cases the complexity of the hydrogen-bondingnetwork required for specificity is increased further by bound water molecules lodged between the protein andthe DNA (Steitz, 1990).Although hydrogen bonding is a very important determinant of molecular complementarity, it isimportant to note that electrostatic interactions can contribute to the specificity of binding (Steitz, 1990).11Double-stranded B-DNA displays sequence-dependent structural polymorphisms, and the subsequentpresentation of phosphate groups does vary in a sequence-dependent manner (Kennard & Hunter, 1989; Somers& Phillips, 1992). The formation of salt-bridges between a sequence-specific DNA-binding protein and itscognate binding site is also influenced by the sequence-dependent deformability of B-DNA (Wu & Crothers,1984), and such deformations can be quite marked (e.g., with EcoR I or CAP; McClann el al., 1986; Schultzet al., 1991). The sequence-dependent structural polymorphism and deformability of B-DNA also help explainhow interactions in the minor groove can contribute to the specificity of binding (Marmorstein et al., 1992).c. The kinetics ofcognate binding-site selection.Sequence-specific DNA-binding proteins typically display remarkable kinetic properties in vitro andin vivo. The maximum rate possible for the bimolecular process of complex formation should in principle occurif this process is diffusion limited, i.e., if during the course of a three-dimensional random walk each collision ofthe protein and its cognate binding site results in complex formation. The Debye-Smoluchowski equation hasbeen used to estimate the diffusion-controlled limit for the rate of complex formation between sequence-specificDNA-binding proteins and their cognate binding sites (von Hippel et al., 1984). In this equation ic and f aredimensionless interaction factors, N0 is Avogadros number, b is the interaction radius (in cm), and Dproteinand Dbinding site are the three-dimensional diffusion constants of the participants (in cm2 sec-1).k a, calc = 4 it 1steric f electrostatic b (Dprotein + Dbinding site ) N0 /1000Many sequence-specific DNA-binding proteins are able to locate their targets at a rate that exceeds thetheoretical diffusion-controlled limit by several orders of magnitude (Lohman, 1986; Record et al., 1991;von Hippel et al., 1984). The ability of sequence-specific DNA-binding proteins to exceed this limit isattributed to their ability to bind to DNA nonspecifically. For statistical reasons nonspecific binding is the mostlikely initial form of association between a sequence-specific DNA-binding protein and DNA. Nonspecificbinding to DNA therefore effectively increases the interaction distance (b in the above equation) for the initialstage of target location. The ability of a nonspecificafly bound protein to subsequently reach its cognate binding12site is believed to be facilitated by intersegment transfer, interdomain association and dissociation, andiniradomain sliding within and between various regions of the genome (von Hippel et al., 1984). During thefinal stage of target location the nonspecifically bound protein is somehow able to track along the DNA until itreaches its cognate binding site. This occurs in Vitro at a rate of up to 1,000 nucleotide pairs per second (for thelac repressor), and during this process sequence-specific DNA-binding proteins are believed to be capable ofrapidly alternating between nonspecific and sequence-specific DNA-binding conformations (Lohman, 1986;Record et al., 1991; von Hippel et al., 1984).d. Cognate binding-site selection in vivo.The study of eukaryotic, sequence-specific DNA-binding proteins fl Vitro has yielded many valuableinsights about how they function. However, the really difficult challenges in this field lie in understanding howthese proteins act on chromatin in vivo. The ionic milieu, DNA topology, and overall structural organization ofthe nucleus is difficult to maintain during subcellular fractionation, and in vitro studies that have ignored thesetechnical difficulties have sometimes yielded results of dubious physiological relevance (see Cook, 1989 for anexcellent discussion of the problem). Most of the studies that have addressed the influence of nucleosome andhigher order chromatin structure on the recognition of specific DNA sequences by regulatory proteins havefocused on “factors” that are involved in the initiation of transcription (in most cases by RNA polymerase II),and the following discussion reflects this bias.The histone octamer consists of two molecules each of the core histones H2A, H2B, H3 and H4, andone molecule of histone Hi is associated with a short region of linker DNA that separates each histone octamer(Felsenfeld, 1992). About 165 bp of DNA is coiled in approximately two superhelical turns around the outsideof the nucleosome core particle. Thus only one side of the DNA helix is presented on the outside of thenucleosome, which may explain why DNA regulatory sequences are often made up of symmetric repeatingunits. Another difference between a DNA target sequence displayed on a nucleosome in vivo and the samesequence studied in Vitro 1S the curvature of the helix axis. The spiraling helix axis of the DNA wound aroundthe outside of the nucleosome differs from the straight helix axis of the short, double-stranded oligonucleotidesthat are used in various in vitro manipulations and most high-resolution structural studies. Supercoiing of DNA13around the histone octamer also causes significant structural distortion of the DNA, with severe kinkingobserved at two positions 1.5 helical turns from the dyad axis of the nucleosome (Hogan et al., 1987).Most investigators agree that the 30-nm chromatin fibre is transcriptionally inert (Amouyal, 1991;Felsenfeld, 1992; Hansen & Ausio, 1992; Kornberg & Lorch, 1991, 1992; Morse, 1992; van Holde et al., 1992).Another point of agreement is that various RNA polymerases are capable of elongating transcripts throughtemplates that contain nucleosomes. However, there appears to be disagreement about whether or notnucleosome formation on gene regulatory sequences represents an accessibility barrier for eukaryotictranscription initiation factors, or whether these same factors actively disrupt nucleosomes and leave the regionsurrounding their cognate binding sites nucleosome-free (Adams & Workman, 1993; Felsenfeld, 1992;Kornberg & Lorch, 1991, 1992; van Holde et at., 1992). The function of topoisomerases, the organization ofDNA into domains consisting of anchored loops, and other aspects of nuclear architecture add further levels ofcomplexity to the problem of how different sequence-specific DNA-binding proteins locate their cognatebinding sites during transcription initiation in vivo (Dworetzky et al., 1992; Getzenberg et al., 1991; Haaf &Schmid, 1991; Stiefet al., 1989).B. REGULATION OF THE TRANSCRIPTION OF EUKARYOTIC HEAT-SHOCK GENES.1. The heat-shock response.The discovery of the heat-shock response is credited to Ritossa’s initial observations of heat-shock-inducible puffing activity, accompanied by the colocalized, ribonuclease-sensitive incorporation of tritiatedcytidine in polytene chromosomes isolated from salivary glands of Drosophila busckii (Ritossa, 1962).Treatment of isolated salivary glands at 25 °C (the normal culture temperature) with either salicylate or2,4-dinitrophenol provoked the same response as a sublethal heat shock (exposure to 30 °C), which showed thata mild temperature elevation is only one of several stress signals capable of inducing the same well-defined,specific transcriptional response (Ritossa, 1962). Considerably more interest in this phenomenon was generatedby a later study in D. melanogaster, which showed that the heat-shock response entails a rapid increase in the14expression of a prominent set of several new proteins, along with an equally dramatic reduction in the synthesisof proteins that were previously expressed under normal conditions (Tissières et at., 1974).Perhaps the most important aspect of the heat-shock response that stimulated subsequent interest wasthe realization that all forms of life appear to share some capacity to mount such a response. Mycoplasma,archaebacteria, eubacteria, fungi, plants, invertebrates, and vertebrates are all capable of responding to a mildtemperature elevation and other stresses by directing the synthesis of a well-defined set of heat-shock proteins(Dascher et a!., 1990; Morimoto et a!., 1990; Trent et at., 1990). The corresponding hsp genes have been foundin nuclear, mitochondrial, and chloroplast genomes (Ashbumer & Bonner, 1979; Hartman et at., 1992;Scaramuzzi et at., 1992).The list of stress signals capable of inducing hsp genes has now expanded to include naturallyoccurring endogenous stimuli (certain hormones, inflammatory response mediators, and other serumcomponents), microbial infection, elevated levels of denatured or improperly folded proteins, anoxia, variousorganic and inorganic chemicals, and temperature elevation (Ananthan et at., 1986; Ashburner & Bonner, 1979;Morimoto et at., 1990). A major question in the molecular biology of the heat-shock response is how thesevarious forms of stress induce the selective transcription of hsp genes.Heat-shock proteins are classified on the basis of their apparent molecular mass (designated inkilodaltons, e.g., HSP7O) and amino acid sequence similarity. The degree of amino acid sequence similaritydisplayed by members of a given class of HSPs can be very high, even when comparing species as different asE. coli and D. melanogaster (e.g., 48% for DnaK and HSP7O; Bardwell & Craig, 1984). In eukaryotes there aremultiple copies of the respective genes encoding members of each class of HSPs (Ashburner & Bonner, 1979).Their homologues in E. coli are single-copy genes that are constituitively expressed at reduced levels undernormal conditions (Georgopoulos et a!., 1990). It was later realized that several eukaryotic HSPs andnoninducible HSP homologues (heat-shock cognate proteins, or HSCs) are also expressed under normalconditions or at certain stages of development (Mg et at., 1991; Morimoto et at., 1990; Schlesinger, 1990).The four major classes of heat-shock proteins are HSP9O, HSP7OIDnaK, HSP6O/GroEL, and lowmolecular-mass (or small) HSPs, where DnaK and GroEL designate E. coli HSPs that were originally defined in15terms of their effects on the growth of bacteriophage lambda (Georgopoulos et al., 1990). The rapid, coordinateexpression of these and other HSPs presumably serves to protect a cell from the deleterious effects of an initial,sublethal heat shock (Hahn & Li, 1990). The expression of HSPs is also believed to confer thermotolerance,which can be defined as the transient protection afforded by a mild heat shock against the deleterious effects of asecond, more severe, and otherwise lethal heat shock (Hahn & Li, 1990; Johnston & Kucey, 1988; but see alsoSmith & Yaffe, 1991 for data challenging this belief). M understanding of how these goals are accomplishedrequires explanations of how various HSPs function.The protective functions of low molecular-mass HSPs are not very well understood. In some casessmall HSPs act as molecular “chaperones’ that stabilize proteins under heat-shock conditions and direct thecorrect refolding of denatured proteins (Jakob et al., 1993). Small liSPs share amino acid sequence similaritywith vertebrate cr-crystaflins, which are found in the lens of the eye (Ingolia & Craig, 1982). Like ct-crystallins,small HSPs are capable of forming very high molecular-mass aggregates in vivo (Ingolia & Craig, 1982).However, it is unclear how these very large aggregates function to protect a cell during heat shock.More is known about the protective functions of the HSP9O, HSP7OIDnaK, and HSP6O/GroELheat-shock proteins (Craig et a!., 1993; Georgopoulos & Welch, 1993; Hendrick & Harti, 1993; Parsell &Lindquist, 1993). Members of the HSP9O class of proteins act as molecular chaperones by forming complexeswith glucocorticoid receptors, proto-oncogene products, various kinases, and other cellular proteins in eukaryoticcells (Ellis & van der Vies, 1991; Schlesinger, 1990). The HSP6O/GroEL class of proteins act as “chaperonins’that direct the folding and assembly of proteins in bacteria (GroEL) or in eukaryotic organelles (Mg et a!.,1991; Cheng et a!., 1989; Schlesinger, 1990). Members of the most prominent class of HSPs, the HSP7OIDnaKfamily, also act as molecular chaperones (Ellis & van der Vies, 1991; Hubbard & Sander, 1991). These proteinsdirect the maturation, unfolding, disaggregation, reactivation, and translocation across biological membranes ofa variety of proteins (Beckman et a!., 1990; Chirico et a!., 1988; Shi & Thomas, 1992; Skowyra et a!., 1990;Yalovsky et a!., 1992). The E. coli DnaK and eukaryotic HSP7O proteins may also be involved in thetranscriptional autoregulation of the heat-shock response (Abravaya et a!., 1992; Baler et a!., 1992; Beckmannet a!., 1992; Gamer et a!., 1992; Liberek & Georgopoulos, 1993; Liberek et a!., 1992; Stone & Craig, 1990).16Considerable attention has been focused on the clinical significance of the expression of heat-shockproteins (Res et al., 1991). Parasites presumably cannot avoid expressing heat-shock proteins, either upontransmission from a poikilothermic vector to a human host, or in response to inflammation, fever, hypoxia,drugs, reactive oxygen intermediates or other stimuli (Mazier & Mattei, 1991). Bacterial infection, especiallyintracellular bacterial infection of human cells (e.g., by mycobacteria), also exposes the infecting microorganismto a variety of stimuli capable of inducing a heat-shock response (Kaufmann, 1991). However, these foreignHSPs contain regions of amino acid sequence similarity with the heat-shock proteins of the host. Furthermore,expression of the latter increases in response to microbial infection, and these self HSPs may be displayed on thesurface of lymphocytes during antigen presentation. The host’s immune system is presented with the verydifficult problem of providing adequate immune protection while avoiding immunopathology.One example of this problem is the ability of a murine anti-HSP7O monoclonal antibody to mediateantibody-dependent cellular cytotoxicity (ADCC) against human H9 lymphoma cells (Di Cesare et at., 1992).Normal H9 cells are not very susceptible to ADCC in the presence of the antibody, but heat shock increases thissensitivity significantly due to increased levels of surface-expressed HSP7O. However, an even greater increasein ADCC was observed in nonshocked H9 cells that were chronically infected with HIV (Di Cesare et al., 1992).Other reports have suggested that inflammatory arthritis, insulin-dependent diabetes, and other autoimmunediseases may be caused by the destruction of host cells bearing epitopes that are cross-reactive with the HSPsexpressed by various infectious agents (Doherty et at., 1991; Life et at., 1991).2. Heat-shock gene promoters.The linear map of a gene designates the nucleotide pair that acts as the transcriptional start site as the+1 position. A designation of “-n” is assigned to a given nucleotide pair in front of or “upstream” from, thetranscriptional start site, and this designation is based on the number (n-i) of nucleotide pairs between the twosites. In eukaryotic protein-coding genes the region of DNA immediately upstream from the transcriptional startsite promotes the formation of a productive transcription initiation complex with RNA polymerase II (pol II).This ‘promoter” region contains short DNA sequence “elements” that act as cognate binding sites forDNA-binding proteins that regulate transcription initiation. This region of DNA is particularly important17because gene expression is frequently regulated at the level of transcription initiation (see Johnson & McKnight,1989; Latchman, 1990; Mitchell & Tjian, 1989).Some promoter elements are relatively common (e.g., the AT-rich sequence or “TATA” element that isfound in most pol II promoters), while others are relatively specific for genes or gene families that arecoordinately regulated at the level of transcriptional initiation. Most of these promoter elements are locatedbetween positions -100 and the start site of transcription. However, pol II transcription is sometimesenhanced—and sometimes dramatically so—by the presence of “enhancers ‘ short DNA sequences that can belocated several thousand base pairs either upstream or downstream from the start site of the gene that theenhancer regulates (see Schena, 1989).Heat-shock gene promoters contain one or more “heat-shock elements”, or HSEs, as well as otherpromoter elements that act in conjunction with HSEs and the TATA element to regulate transcription initiation(Petham, 1982; see also Nover, 1987). HSEs can also be viewed as heat-shock-inducible enhancers, and theycan function at considerable distances from the transcription start site (Bienz & Petham, 1987a). The followingcriteria were used to identify the original HSE consensus sequence: HSE-like sequences were present in one ormore copies in all known heat-shock gene promoters; the deletion of HSE-like sequences from the promoter of aknown heat-shock gene abolished the heat-inducibility of that gene; and the introduction of an HSE consensussequence in the promoter of a noninducible reporter gene rendered the reporter gene heat-inducible (Petham,1982; Pethain & Bienz, 1982). The conserved nature of the HSE was subsequently confirmed by a large numberof gene-transfer studies showing that heterologous heat-shock genes can only be expressed at the temperaturerequired to induce the host’s heat-shock response (see Nover, 1987).The original consensus sequence for the HSE was defined as J G G A A T N T T C T A.i A(Petham, 1982). However, later studies showed that a more precise defmition of the HSE consensus sequence ismultiple consecutive repeats of the sequence N G A A N arranged in alternating orientation, with multiples ofthe trimeric structure N G A A N N T T C N N G A A N representing an optimal variation thereof (Aminet al., 1988; Perisic et at., 1989; Xiao & Lis, 1988; but see also Raibaud, 1990). Within each N G A A N18pentamer there is a strong preference for a deoxyadenylate residue at position 1, and yet another redefinition ofthe HSE consensus sequence as A G A A N has recently been proposed (Femandes et at., 1994).The HSE has been shown to serve as the cognate binding site for a sequence-specific DNA-bindingprotein, and in some cases this protein has been shown to activate hsp gene-specific transcription in vitro(Parker & Topol, 1984; Wu, 1984a, 1984b, 1985). The consecutive repeat structure of the HSE suggested thatthis heat-shock gene transcription factor (HSTF or HSFt) binds to the HSE as a multimer, and this predictionproved correct (Shuey & Parker, 1986a; Sorger & Petham, 1987; Topol et al., 1985).An important question in the study of gene expression is determining what distinguishestranscriptionally active from inactive chromatin. The reversibility, simple induction mechanism, and highlyconserved nature of the transcriptional response of heat-shock genes have prompted several studies of thechromatin structure surrounding these loci. Most of these studies have focused on the detection of DNase Ihypersensitive sites in heat-shock gene promoter regions, and the association of topoisomerases with transcribedheat-shock genes (e.g., Costlow & Lis, 1984; Fleischmann et at., 1984; Gilmour et at., 1986; Kroeger & Rowe,1992; Rowe et at., 1986; Udvardy & Schedi, 1993; Wu, 1980).Another series of studies have examined the association of Drosophila RNA polymerase II with anhsp7O heat-shock promoter in vivo. As expected, a high density of pot II molecules were observed along theentire length of the Drosophila hsp7O gene after induction of the heat-shock response (Gilmour & Lis, 1986).However, even in the absence of heat shock, approximately one molecule of stalled RNA polymerase II, towhich a 20- to 30-ribonucleotide long hsp7O gene transcript was attached, was observed to be transcriptionallyengaged between positions -12 and +65 on each hsp7O promoter (Gilmour & Lis, 1986; Lee et at., 1992; O’Brien& Lis, 1991; Rougvie & Lis, 1988).These findings help explain the rapid nature of the transcriptional response during heat shock, and theyimply that the transcriptional activation domain of Drosophita HSF can directly or indirectly alleviate thist There is no standard nomenclature for transcription factors. Presumably the initially used designation, HSTF,was meant to refer to HSE-specific DNA-binding proteins that had been shown to function as activators of lispgene transcription (either in vitro or in vivo). The more commonly used term HSF (heat-shock factor) nowrefers to any HSE-specific DNA-binding protein, even if this protein’s ability to function as an hsp gene-specifictranscriptional activator has yet to be demonstrated (e.g., Czarnecka et a!., 1990; Gallo et at., 1991; Hensoldet al., 1990; Sarge et at., 1991; Scharf et al., 1990).19transcriptional arrest mechanism (Gilmour & Lis, 1986). Thus Drosophila HSF appears to function as atranscription elongation factor; i.e., it acts at a step after the initiation of transcription (Lis & Wu, 1993;Rasmussen & Lis, 1993). It is less clear whether Drosophila HSF also acts as a transcription initiation factorafter the initial round of hsp gene transcription has been allowed to proceed unhindered. Furthermore, it isunknown if RNA polymerase II initiates the transcription of heat-shock genes prior to heat shock in othereukaryotic organisms.3. Heat-shock transcription factor (HSF).a. Interspecies and intraspecies diversity.In eukaryotes both the heat-shock response and the multimeric, pentanucleotide-repeat structure of theHSE are highly conserved. In addition, some of the structural features and functional attributes of HSF arecommon to all known isolates of this protein. However, the HSFs isolated from different eukaryotic speciesdisplay remarkably little conservation of their amino acid sequences, show limited serological cross-reactivity,and show significant differences in the mechanistic details of their DNA-binding and transcriptional activationfunctions (see Sorger, 1991; Zimarino et al., 1990a). The relative molecular mass of the 11SF polypeptide(predicted from the DNA sequence of the corresponding hsf gene) ranges from a high of 93,000 (inS. cerevisiae; Sorger & Petham, 1987) to a low of 35,000 (for an HSF in tomato; Scharf et at., 1990). Finally, inaddition to this interspecies diversity, the human genome contains more than one hsf gene, and significantintraspecies diversity is exhibited by these genes (Rabindran et al., 1991; Schuetz et at., 1991). Intraspeciesdiversity is also displayed by the multiple nsfgenes found in the chicken, mouse, and tomato genomes (Nakai &Morimoto, 1993; Sarge et at., 1991; Scharfet at., 1990).b. Quaternary structure.The quaternary structure of 11SF is a homotrimer and multiples thereof, which form in solution and onHSE-containing DNA sequences (Perisic et at., 1989; Sorger & Nelson, 1989). Binding of individualhomotrimer subunits on HSE-DNA is highly cooperative, and each 11SF monomer is believed to contact oneN G A A N pentanucleotide sequence (Shuey & Parker, 1986a; Topol et at., 1985; Xiao et a!., 1991). Stable20binding of the HSF trimer requires that at least two of the three monomers contact consecutive N G A A Nsequences, which can be arranged in any orientation (Pensic et at., 1989). However, in the absence of a thirdconsecutive HSE pentanucleotide sequence, the binding of the third HSF monomer may not occur, or it mayoccur at a third, appropriately spaced N G A A N sequence that is separated from the other twopentanucleotide sequences (see Sorger, 1991). The binding of HSF to the HSE introduces a bend in the DNAin vitro, and causes various DNA helix distortions in vivo (Gross et a!., 1990; Shuey & Parker, 1986b). Thebinding of HSF trimers on DNA sequences containing multiple HSEs is also highly cooperative, and ahexaineric HSF complex may be a functionally significant structure in vivo (Cbs et at., 1990; Xiao et at.,1991).c. General properties of the HSF monomer.The tertiary structure of the HSF monomer contains domains for transcriptional activation,DNA-binding, and HSF trimerization. There is also a regulatory “masking” region whose three-dimensionalstructure appears to involve several parts of HSF (Cbs et al., 1990; Nieto-Sotebo et at., 1990; Sorger, 1990).HSF is an acidic protein overall, with a basic region covering the N-terminal third of the protein and an acidicregion extending over the rest of the protein (Nieto-Sotelo et at., 1990). 11SF migrates anomalously onSDS -PAGE gels, yielding an apparent molecular-mass value significantly larger than the one which would bepredicted from the DNA sequence of the corresponding hsf gene (e.g., 150 vs. 93 kDa; Sorger & Pelham, 1988).HSF is a phosphoprotein in some species (Chang et at., 1993; Larson et at., 1988; Mivechi et at., 1994; Sargeet at., 1993a, 1993b), and in budding yeast increased phosphorylation is believed to be either a prerequisite for,or consequence of derepression of the transcriptional activation domain during heat-shock (Sorger & Petham,1988).d. The trimerization domain.The HSF trimerization domain is located near the centre of the protein’s primary structure (Cbs et at.,1990; Sorger, 1990; Sorger & Nelson, 1989). This domain contains a highly conserved region of cx-helicalsecondary structure that is distinguished by the occurrence of hydrophobic amino acids every seven residues, or21every two turns of the a -helix. These “hydrophobic heptad repeats” leave an uninterrupted string ofhydrophobic side chains exposed along one face of the helix. In most cases the trimerization domain is made upof one longer a-helix that is followed by two shorter a-helices that are one amino acid out of phase with theheptad repeats of the first a-helix (Cbs et a!., 1990). The trimerization domain cc-helices are amphipathic, andamino acids bearing charged side chains are aligned on the face of the helix opposite from the hydrophobicheptad repeats. Deletion studies have shown that these cc-helical regions are required for trimerization (Sorger,1990; Sorger & Nelson, 1989).It has been hypothesized that the trimerization helices of three HSF monomers form a parallel,three-stranded coiled coil that gives the resulting HSF trimer its structural integrity and DNA-binding flexibility(Sorger & Nelson, 1989; Peteranderl & Nelson, 1992). In such a structure the three right-handed cc-helices windaround each other in a left-handed supercoil (Cohen & Parry, 1990). The formation of this supercoiled structureis dependent on interchain hydrophobic interactions involving nonpolar amino acid side chains located on theappropriate faces of the three cx-helices. This trimerization mechanism is somewhat analogous to thedimerization mechanism mediated by the two parallel a-helices that form the “leucine zipper” structure found insome eukaryotic transcription factors (e.g., C/EBP; Landschulz et a!., 1988). The coiled-coil structure proposedfor HSF is also analogous to the three-stranded coiled coil formed by influenza virus hemagglutinin, and it isnoteworthy that the latter is capable of a functionally significant conformational change under certain conditions(i.e., at low pH; Skehel et a!., 1982).e. The DNA-binding domain.The HSF trimerization and DNA-binding domains are the only two regions where significant aminoacid sequence similarities are displayed by HSF isolates (Cbs et a!., 1990; Nieto-Sotebo et a!., 1990; Rabindranet al., 1991; Sarge et a!., 1991; Scharf et a!., 1990; Schuetz et a!., 1991; Sorger, 1990). The DNA-bindingdomain is located between the N-terminus and the central trimerization domain of the protein’s primarystructure. A noteworthy feature of the DNA-binding domain is the presence of two short cz-helices that aresimilar in amino acid sequence to the recognition helix of bacterial a factors (Cbs et a!., 1990; Helmann &Chamberlin, 1988). It has also been shown that a yeast gene involved in the suppression of flocculation (sf1)22contains a region encoding a stretch of 108 amino acids that displays 44% identity with a region in theDNA-binding domain of HSF (Fujita et al., 1989). In addition, a multicopy suppressor (SKN-7) of a mutationaffecting cell-wall f3-glucan assembly in yeast contains a 26 amino acid region that displays 81% identity with aregion in the DNA-binding domain of HSF (Brown et al., 1993).The DNA-binding domain of K lactis HSF has been crystallized and the crystal structure determined at1.8 A resolution by X-ray diffraction methods (Harrison et al., 1994). The three cx-helices of the HSFDNA-binding domain form a compact bundle capped by a four-stranded antiparallel 13-sheet, with anal-131-f32-a2-cL3-133-134 sequence of secondary-structure elements (Harrison et al., 1994). The a2 and a3helices form a helix-turn-helix DNA-binding motif. The first a-helix of this motif in HSF (a2) contains ana-helical bulge and a proline-induced kink, and this proline residue (amino acid 238 in K. lactis HSF) isinvariant in HSF isolates (Harrison et al., 1994). This proline residue is also found at the appropriate position inthe region of sequence identity of SKN-7 (but not SFL-1) (Harrison et al., 1994).The solution structure of the DNA-binding domain of Drosophila HSF has been studied by NMRmethods, and these results agree with the X-ray structural data mentioned above (Vuister et al., 1994). Thisstudy also examined the binding of15N-labeled HSF to a short DNA duplex bearing one copy of the HSE, andconcluded that the third a-helix is the recognition helix. The overall structure of the helix-turn-helixDNA-binding motif of HSF, the prokaryotic catabolite activator protein (CAP), and members of theIINF-3lforkhead family of eukaryotic transcription factors are similar (Harrison et al., 1994; Vuister et al.,1994).f The transcriptional activation domains.The transcriptional activation domains of various HSF isolates do not display any significant aminoacid sequence similarity (Cbs et al., 1990; Nieto-Sotelo et al., 1990; Rabindran et al., 1991; Sarge et al., 1991;Scharf et a!., 1990; Schuetz et a!., 1991; Sorger, 1990). In addition, different researchers have used differenttypes of heat-shock promoters to define different types of transcriptional activation domains along the primarystructure of HSF. Another complication that clouds the distinction between different types of transcriptionalactivation domains is the function of the “masking” region, a region which presumably acts as a regulatory23interface between HSF and another protein, possibly an HSF repressor. These difficulties have led to two verydifferent maps of the location of different types of transcriptional activation domains along the primary structureof yeast HSF (Nieto-Sotelo et al., 1990; Sorger, 1990).Sorger (1990) has characterized two separate transcriptional activation domains, one “transient” and theother “sustained”, at the N-terminal and C-terminal regions, respectively, of HSF from the budding yeastS. cerevisiae. These two domains are responsible for the inducible and constitutive transcriptional activationcapabilities of yeast HSF. The former capability is not detectable in the absence of heat shock, whereas thelatter capability is specific for certain heat-shock promoters and is detectable at different levels at differentgrowth temperatures.Transient and sustained transcriptional activation domains have also been distinguished inKluyveromyces lactis HSF, but not in the HSFs isolated from higher eukaryotes (Chen et al., 1993). Thetranscriptional activation domain that has been mapped in the HSFs isolated from higher organisms is near theC-terminus of the protein (see Sorger, 1991). Furthermore, the distinction between transient and sustainedtranscriptional activation domains in yeast 11SF is not straightforward. Deletion of the C-terminal, “sustained”transcriptional activation domain from the hsf gene in S. cerevisiae only affects viability at temperatures abovethe threshold for hsp gene expression (Sorger, 1990). However, yeast 11SF is essential for survival at all growthtemperatures, i.e., both below and above the temperature threshold for hsp gene expression (Gab et al., 1993;Sorger & Petham, 1988; Wiederrecht et al., 1988).g. The regulatory “masking” region.A study of the HSFs isolated from two species of budding yeast have been used to determine some ofthe amino acid sequences that make up the masking region (Jakobsen & Petham, 1991). The component parts ofthis region appear to include some of the conserved amino acid sequences located in the DNA-binding andHSF-trimerization domains, as well as a conserved heptapeptide, RXLLKNR, located adjacent to the C-terminaltranscriptional activation domain. Deletion of this conserved, heptapeptide sequence element (CE2) resulted inthe constitutive expression of a heat-shock regulated reporter gene at a level that is usually only observed afterheat shock.24Chimeric genes that encode various regions of HSF joined to the transcriptional activation domain ofthe VP16 protein of herpes simplex virus have also been used to study the masking region (Bonner et a!., 1992).The masking region of HSF was able to correctly regulate this heterologous transcriptional activation domain.Weak point mutations affecting CE2, or much stronger point mutations affecting a methionine residue (met 232)in the DNA-binding domain, resulted in the constitutive derepression of yeast HSF-VP16 chimeras.Methionine 232 is very close to the N-terminal boundary (between amino acids 208 to 222) of the yeastHSF sequences that are involved in the masking region (Nieto-Sotelo et a!., 1990). This boundary was alsodetermined using gene fusions, but in this case a gene fragment encoding a heterologous DNA-binding domainwas joined to various regions of the yeast lisf gene. The masking region of HSF was able to correctly regulate achimera that had the DNA-binding specificity of the yeast transcription factor AP-1, but which retained part ofthe DNA-binding domain and the entire C-terminal transcriptional activation domain of HSF. However, itremains unclear how methionine 232, the CE2 heptapeptide, other parts of HSF, and possibly other proteins areinvolved in masking the C-terminal transcriptional activation domain at temperatures below the threshold forlisp gene expression.4. The biochemical mobilization and demobilization of HSF.a. Signal transduction.The regulation of the expression of a given heat-shock gene during the heat-shock response depends onthe lisp gene in question, the species and physiological status of the organism or cell, and the nature and durationof the stimulus. Most lisp genes do not contain introns, but in the rare cases where they do occur, expression ofsuch genes may be regulated at the level of RNA splicing during heat shock (see Yost et a!., 1990a).Translational regulation of the heat-shock response occurs in some organisms, or in others at certain stages ofdevelopment (see Yost et a!., 1990b). In these cases hsp mRNA is selectively translated during heat shock,while the translation of non-heat-shock mRNA is arrested until after the inducing stimulus has been withdrawn.A somewhat related form of RNA processing that can affect lisp gene expression is lisp mRNA-specificdegradation during recovery (DiDomenico et al., 1982; see Yost et a!., 1990a). However, the transcriptional25regulation of heat-shock gene expression is particularly important. Most of the diverse stimuli capable ofinducing the heat-shock response are believed to act directly or indirectly to influence either the binding of HSFto the HSE, the subsequent unmasking of the inducible transcriptional activation domain of HSE-bound HSF, orboth of these events.Many of the various stimuli capable of inducing the heat-shock response are also able to compromisethe proper folding and assembly of proteins. A variety of evidence supports a role for the intracellularaccumulation of denatured proteins as a common signal for inducing the transcription of heat-shock genes (e.g.,Ananthan et al., 1986; Edington et al., 1989). One hypothesis that was based on these observations suggestedthat HSF is a target for ubiquitin-mediated degradation under normal conditions (Bienz & Petham, 1987b).However, under heat-shock conditions HSF was thought to escape this fate due to the accumulation of denaturedproteins to levels that overwhelm the protein-degrading capacity of the cell. This hypothesis is consistent withthe known susceptibility of HSF to protease activity in Vitro (Sorger & Petham, 1987; Wiederrecht et al., 1987;Wu et al., 1987).The results of experiments carried out using the protein synthesis inhibitor cycloheximide did notsupport the ubiquitin-mediated, HSF turnover hypothesis. Addition of this drug during a heat-shock inductionprotocol that involved alternating periods of induction followed by recovery did not prevent the heat-shockresponse from occurring during the initial or subsequent periods of induction (Zimarino & Wu, 1987; Zimarinoet al., 1990a). These results showed that (i) HSF exists in cells prior to heat shock, (ii) the concentration of HSFdoes not increase during heat shock, (iii) HSF activity is not regulated by degradation, and (iv) HSF appearsinstead to undergo some type of reversible change that allows it to function during heat shock, return to itspreexisting state after withdrawal of the inducing stimulus, and then be able to repeat the process if exposed toanother heat shock.Studies of glutamate and glutamine metabolism in normal and heat-shocked Drosophila cells has led toa series of interesting findings on the role of glutamine in supporting high-level expression of HSPs during heatshock (Cai et al., 1991; Sanders & Kon, 1991, 1992). The following cell culture conditions were found tosupport maximal HSP synthesis under heat-shock conditions: (I) prior addition of glutainine, (ii) prior addition26of a glutamine analogue, (lii) stimulation of endogenous glutamine synthesis by the prior addition of glutamateand NH, and (iv) stimulation of endogenous glutainine synthesis by the prior addition of glutamate and analanine aminotransferase inhibitor, with the latter compound acting to increase intracellular NH4 levels.However, it is unclear from these reports exactly how intracellular glutamine levels affect HSF activity duringheat shock.b. The response ofHSF to heat shock.The response of HSF to heat shock has been studied extensively in tissue-culture cells and salivarygland squashes from the fruit fly Drosophila melanogaster (Westwood et al., 1991). dmHSFt is already presentin the nucleus prior to heat shock, and it appears to be either an inactive homodimer (Mr of 220,000) or in aninactive complex with some other protein or proteins. dmHSF responds to heat shock by rapidly associatingwith a large number of chromosomal loci. In salivary gland squashes the number of polytene sites to whichdmHSF binds during heat shock exceeds the number of known heat-shock genes, and many of the former havebeen identified as key developmental loci. These unexpected results suggest that HSF may act as an induciblepleiolropic repressor in addition to its function as an hsp gene-specific transcriptional activator (Westwood et a!.,1991).Heat shock also transforms the inactive, 220 kDa form of dmHSF into a high-Me oligomer that is nowcapable of binding to the HSE (Westwood et a!., 1991). The chromatin structure surrounding heat-shockpromoters is believed to be maintained in a nucleosome-free state, and binding of dmHSF to HSEs on achromatin template requires the presence of the general transcription factor TFIID (Lis & Wu, 1993; Tayloret a!., 1991). After dmHSF binds to the HSE the transcriptional activation domain is unmasked, thetranscriptional arrest of RNA polymerase II is relieved, and robust hsp gene transcription proceeds unhindered(Gilmour & Lis, 1986; Lee et a!., 1992; Rougvie & Lis, 1988).The response of HSF to heat shock has also been studied extensively in the budding yeastSaccharomyces cerevisiae. scHSF is also present in the nucleus under normal conditions, but it is already boundt In this thesis the first and second lower case letters in abbreviations such as dinHSF refer to the genus and thespecies, respectively.27to the HSE prior to heat shock (Sorger et al., 1987). Heat shock results in the unmasking of the scHSFtranscriptional activation domain, which is accompanied by an increase in the extent of scHSF phosphorylation(Sorger & Petham, 1988). The response of 11SF to heat shock in the budding yeast Kluyveromyces lactisappears to be similar, and the Kluyveromyces hsf gene can replace the essential hsfgene of S. cerevisiae (Chenet al., 1993; Jakobsen & Petham, 1991). However, two reports disagree about the ability of HSF to bind to theHSE prior to heat shock in the fission yeast Schizosaccharomyces pombe (Gallo et al., 1991; Zimarino et al.,1990a).The response of HSF to heat shock in mouse cells is quite complicated. The behaviour of an HSFactivity in mouse fibroblasts appears to follow the pattern of heat-inducible, HSE-specific DNA-bindingdisplayed by HSF in other higher eukaryotes (Mezger et al., 1989; Sarge et a!., 1993a, 1993b). However, thebehaviour of the mmHSF activity present in undifferentiated murine embryonal carcinoma (EC) cell lines is veryunusual (Mezger et a!., 1989). An mmHSF activity is readily detectable prior to heat-shock in all EC cell linesexamined, and under normal conditions all of these cell lines constituitively express a well-defined subset ofHSPs. This mmHSF activity increases upon heat shock in EC cell lines that are capable of the inducibleexpression of HSPs (i.e., the complete set of HSP5), while the same activity decreases upon heat shock in thoseEC cell lines that are incapable of the inducible expression of HSPs. However, if the uninducible EC cell linesundergo differentiation in vitro, they then exhibit the same mmHSF activity and heat-shock response as theinducible EC cell lines (Mezger et a!., 1989).Although it is frequently stated that HSFs from higher eukaryotes and budding yeast differfundamentally in their ability to bind DNA prior to heat shock, the distinction does not always hold. Some formof HSF binding to the HSE, or the binding of some HSF isoform, has been reported to occur prior to heat shockin Drosophila Kc cells (but not SL2 cells), human HeLa cells, and soybeans (Czarnecka et a!., 1990; Mosseret a!., 1988; Parker & Topol, 1984). However, in some cases this activity may be an artifact of the method usedto prepare the control extract used in the EMSA binding reactions (inadvertent induction), and in other cases theconstitutive HSF activity appears to be distinct from the activity observed after heat shock. These discrepancies28may also be explained by the presence of more than one hsf gene in some species, which raises the theoreticalpossibility of heteromultimer formation (Rabindran et at., 1991; Schuetz et at., 1991).The binding of HSF to the HSE is therefore necessary but not sufficient for the transcriptionalactivation of heat-shock genes. Further evidence to support this view is the ability of salicylate to induce anHSF activity in human HeLa cells, but its inability to induce the transcription of heat-shock genes (Jurivichet at., 1992). In addition, murine erythroleukemia (MEL) cells appear to express some form of constitutive HSFactivity, despite their inability to transcribe lisp genes at any growth temperature (Hensold et at., 1990).The HSF present in the nuclei of nonshocked Drosophila cells is rapidly lost to the cytosol duringsubcellular fractionation (Westwood et at., 1991). This nominally cytosolic HSF does not display theHSE-specific DNA-binding activity that can be detected in nuclear extracts prepared from heat-shocked cells(Zimarino & Wu, 1987). However, dmHSF isolated from nonshocked cells is capable of HSE-specificDNA-binding in vitro if it is exposed to either low pH or a temperature which induces heat shock in vivo, or if itis incubated with an antibody that is specific for the dmHSF present in heat-shocked cells (Zimarino el at.,1990b). Nominally cytosolic HSF obtained from nonshocked human HeLa cells is also incapable ofHSE-specific binding in vitro, but it is able to do so if it is exposed to low pH, calcium ions, nonionic detergents,chaotropes such as urea, or a temperature which induces heat shock in vivo (Mosser et at., 1990).The genes encoding several HSFs have been cloned and the functions of their expression productsstudied in detail both in vivo and in vitro. Two genes encoding HSF proteins (proteins designated mHSF1 andmHSF2; herein refeffed to as mmHSF1 and mmHSF2) have been cloned from the mouse genome (Sarge et a!.,1991). These genes were expressed using an in vitro translation system and the response of the two mmHSFs toheat shock was studied. mmHSF2 can only bind to the HSE prior to heat shock, but not after, while the moreabundant mmHSF1 can only bind to the HSE after heat shock. However, expression of the mmhsj’2 gene isundetectable, and mmHSF1 messenger RNA can only be detected in certain cell types (Surge et a!., 1991).The results of most hsf gene expression studies support the contention that HSF has an intrinsiccapability for HSE-specific DNA-binding that is expressed at all growth temperatures in budding yeast, butwhich is frequently, but not always, under negative control in nonshocked cells of higher eukaryotes (Jakobsen29& Petham, 1988; Sorger et al., 1987). Thus when the Drosophila hsf gene is expressed in E. coli, therecombinant HSF is quite active as an HSE-specific DNA-binding protein without any heat treatment (Cbset al., 1990). However, if the same gene is expressed in Xenopus oocytes, the HSF produced under nonshockconditions is inactive as an HSE-specific DNA-binding protein unless it is heated in vitro (Cbs et al., 1990).These gene expression studies suggest that in vivo HSF may either undergo a conformational change,bind to or act as a substrate for one or more proteins, or that some combination of these events may occur.These events presumably act in a reversible fashion to regulate the ability of HSF to bind to the HSE, or theability of HSE-bound HSF to transcriptionally activate hsp gene expression, depending on the species. Thein vitro activation of the HSE-specific DNA-binding function of uninduced, metazoan HSFs by exposure to heat,low pH, calcium ions, nonionic detergents, or chaotropes has been explained by the ability of such agents todisrupt the interaction between HSF and a putative repressor, or to cause a transition from an inactive to anactive HSF conformation (Mosser et al., 1990). However, it appears that increases in intracellular hydrogen-ionor calcium-ion concentration are only capable of influencing the HSE-specific DNA-binding activity of HSF.During heat shock in Drosophila cells the intracellular pH drops (from 7.4 to 6.9) and the concentration ofcalcium rises (from 200 to 1900 nM), but these changes are neither sufficient nor required for induction of theheat-shock response (Drummond et al., 1986).Gene expression studies have also addressed the role of the trimerization domain of HSF. If this regionof K lactis HSF is deleted, transformed yeast carrying the disrupted gene are viable but hsp gene transcription isconstitutive (Chen et al., 1993; Jakobsen & Petham, 1991). This result is very surprising given the conservednature of the domain and the in vitro evidence that HSF binds to the HSE as a thmer or multiples thereof. Inbudding yeast the HSF-irimerization domain may only be required to mask the transcriptional activation domainat temperatures below the threshold for hsp gene expression.The hsf genes of Drosophila melanogaster, Schizosaccharomyces pombe, Homo sapiens, Mus muscus,and Gallus domesticus all encode a fourth hydrophobic heptad repeat between the trimerization domain and theC-terminal transcriptional activation domain (Cbs et al., 1990; Gallo et al., 1993; Nakai & Morimoto, 1993;Rabindran et al., 1991; Sarge et al., 1991; Schuetz et al., 1991). This fourth hydrophobic heptad repeat is not30encoded by the hsf genes of budding yeasts. These observations have led to the hypothesis that in nonshockedS. pombe and metazoan cells this fourth amphipathic a-helix inactivates HSF via intramolecular hydrophobicinteractions with the trimerization domain. This would inhibit the intermolecular hydrophobic interactionsrequired for HSF oligomerization and HSE-specific DNA-binding (Rabindran et al., 1993; Lis & Wu, 1993;Westwood & Wu, 1993).This hypothesis appears to be inconsistent with the results of gene expression studies carried out inE. coli with the Drosophila and later the human hsf genes (Cbs et al., 1990; Rabindran et at., 1991; Schuetzet at., 1991). The expression products of these genes are quite capable of HSE-specific DNA-binding in theabsence of any heat treatment. However, the formation of an inactive HSF monomer in nonshocked S. pombeand metazoan cells may involve a particular HSF conformation and/or some other cellular protein to stabilizesuch a structure in vivo.Perhaps the most puzzling results of hsf gene expression studies concern the role phosphorylation playsin the regulation of HSF-mediated transcriptional activation (Mivechi et at., 1994). Murine HSF1 isphosphorylated in response to heat shock, but this event appears to have no functional significance (Sarge et at.,1993a, 1993b). However, when a human hsp7O promoter-linked reporter gene was expressed in murine N-18neuroblastoma cells, the inhibition of protein phosphatase 2A by okadaic acid (an inhibitor of serine/threoninespecific phosphatases) increased the expression of the reporter gene under heat-shock conditions (Chang et at.,1993). Similar results were obtained with Chinese hamster HA-i cells, but when sodium vanadate (an inhibitorof tyrosine-specific phosphatases) was added to the culture medium, heat-shock gene expression decreased(Mivechi et at., 1994).An HSF activity present in HeLa cells also appears to be phosphorylated in response to heat shock(Larson et at., 1988). However, when either of the two human hsf genes are expressed in E. coli, the proteinproducts of these genes are capable of hsp gene-specific transcriptional activation in vitro without any heattreatment (Rabindran et at., 1991; Schuetz et al., 1991). There is no evidence that dmHSF is phosphorylatedbefore or after heat shock, and bacterially expressed dmHSF is also capable of hsp gene-specific transcriptionalactivation in vitro without any heat treatment (Cbs et at., 1990; Wu et at., 1987). It is therefore unclear if31increased 11SF phosphorylation in vivo is a prerequisite for, or a consequence of, the derepression of itstranscriptional activation domain.c. An autoregulatory role for heat-shock proteins?The expression products of the hsp7O gene family are the most abundant heat-shock-inducible proteinsin eukaryotic organisms. When Drosophila cells are heat-shocked the expression of hsp7O is quantitativelyrelated to the degree of stress (DiDomenico et al., 1982). Furthermore, as the concentration of HSP7O increasesduring heat shock its expression is self-regulated at two levels: by the repression of hsp7O transcription and thedestabilization of hsp7O mRNA. Exposure of Drosophila cells to amino acid analogues during heat shockprevents the synthesis of functional HSPs. Under these conditions the self-regulation of hsp7O mRNA synthesisand stability is abrogated, resulting in the accumulation of extremely large amounts of hsp7O mRNA(DiDomenico et al., 1982).Two related discoveries renewed interest in the possibility of an autoregulatory role for HSP7O in thetranscriptional regulation of the heat-shock response (Craig & Gross, 1991; Stone & Craig, 1990). Thediscovery that HSP70 acts as a molecular chaperone provided a theoretical basis for the interaction of HSP7Owith some component of the transcriptional apparatus of the cell. The second discovery that suggested anautoregulatory role for HSP7O was the finding that DnaK, the E. coli homologue of HSP7O, acts in an indirectway to regulate the heat-shock response at the transcriptional level.The E. coli “core” RNA polymerase (designated as E) requires a subunit (designated as a70, where thesuperscript refers to the subunit’s Mr x 10-s) to form a holoenzyme (Ea70) that is capable of recognizing thepromoter region of most genes (Helmann & Chamberlin, 1988). However, recognition of heat-shock promotersrequires the binding of an alternate sigma factor (a32) to core RNA polymerase (Grossman et al., 1984).Sigma-32 is very unstable, and during heat shock both the amount and stability of a32 increase (Straus et al.,1987, 1989). Sigma-32 is therefore both a heat-shock protein and a regulator of heat-shock gene transcription.Transcription of rpoH, the gene that encodes a32, can be initiated at one of four promoters (Erickson & Gross,1989; Nagai et al., 1990; Wang & Kaguni, 1989). The predominant RNA polymerase holoenzyme (Ea70) isrequired by three of these promoters. However, at high growth temperatures a second alternate sigma factor32(0E) is required for the expression ofa32 and a limited number of other heat-shock proteins (Erickson & Gross,1989; Wang & Kaguni, 1989).Three E. coli heat-shock proteins—DnaJ, DnaK, and GrpE—have been shown to form complexes witha32 (Gamer et at., 1992; Liberek & Georgopoulos, 1993; Liberek et at., 1992). The complex formed betweenDnaJ and a32 is unaffected by ATP, whereas the DnaK-a32and GrpE-a32complexes can be disrupted by ATP(Gamer et at., 1992). DnaK and HSP7O contain a weak ATPase activity, and the requirement for ATPhydrolysis to effect the disruption of protein-protein complexes is an integral feature of chaperone activity (seeMg et at., 1991). Evidence supporting an autoregulatory role for DnaK during the heat-shock response has alsocome from a study of bacteria that carry the dnaK756 mutation. These bacteria produce a form of DnaK that isunable to associate with a32, and in this strain heat-shock proteins are overexpressed at all growth temperatures(Liberek et at., 1992). The association of HSPs and a32 appears to be an essential feature of the transcriptionalautoregulation of the E. coli heat-shock response.Direct evidence for an association between HSP7O and HSF has also been reported (Abravaya et at.,1992; Baler et at., 1992; Beckmann et at., 1992). In vitro studies showed that in HeLa cells HSF forms acomplex with HSP7O that cannot be dissociated by ATP (Baler et at., 1992). No complexes were observed toform between HSP9O and HSF or between HSP7O and the transcription factors 0Th or Spl (Baler et at., 1992;but see Nadeau et at., 1993 for a report of HSP9O-HSF complex formation). However, the role of ATP appearsto be in dispute, as an HSP7O-HSF complex that can be dissociated by ATP has been detected in human K562cells (Abravaya et at., 1992). When these experiments were carried out with HeLa cell extracts as a source ofHSF the addition of ATP increased the stability of the HSF-HSP7O complex, and this was attributed to therelease of HSP7O from other cellular proteins. Although the HSF-HSP7O complex detectable in human HeLa orK562 cells is still capable of HSE-specific DNA-binding, it is not known if formation of the complex preventsthe transcriptional activation of heat-shock genes.In vivo studies showed that if the protein synthesis inhibitor cycloheximide is added to HeLa cells priorto the administration of a moderate heat shock, both the detectable level of HSF (as an HSE-specificDNA-binding activity) and the magnitude of the heat-shock response are much less than what would be33observed in the absence of prior exposure to cycloheximide (Baler et aL, 1992; see also Zimarino et al., 1990a).Under these conditions the HSP7O present in the cell prior to heat shock (i.e., due to basal synthesis) isunderutilized due to the inadequate demand for its chaperoning services during the folding of newly synthesizedproteins. The ratio of free HSP7O to HSF is high, and the subsequent exposure of the cells to a mild heat shockdoes not result in the accumulation of enough denatured proteins to exhaust the available supply of HSP7O.Hence HSF remains complexed with HSP7O and a very weak heat-shock response is observed (Baler et aL,1992).A similar phenomenon is observed when chicken embryo cells are cultured in the presence of proteinkosmotropest such as glycerol or deuterium oxide (Edington et al., 1989). The presence of the kosmotropepresumably reduces the demand for the chaperoning activity of HSP7O, and the fraction of HSF complexed withHSP7O remains high. Under these conditions the response to a mild heat shock is much less than what isobserved in the absence of the kosmoirope (Edington et al., 1989).Exposure of HeLa cells to cycloheximide prior to the administration of a severe heat shock does notcause any decrease in the level of HSF or the magnitude of the heat-shock response that is nonnally observedunder these conditions in the absence of prior exposure to cycloheximide (Baler et al., 1992). Under theseconditions the chaperoning function of HSP7O is in high demand due to the rapid accumulation of large amountsof denatured proteins, the fraction of HSF complexed with HSP7O decreases, and a vigorous heat-shockresponse is observed. Excessive demand for the chaperoning function of HSP7O can also be stimulated atnormal growth temperatures by exposure of cells to a variety of agents (i.e., heat-shock inducers) which causethe denaturation of cellular proteins. This can be accomplished directly by the micro-injection of denaturedproteins, but not their properly folded counterparts (Ananthan et al., 1986).There is other evidence to support the hypothesis of HSP7O-mediated autoregulation of the heat-shockresponse. The forced overexpression of HSP7O in transfected human cells reduces the extent of HSF activationafter heat shock (Mosser et al., 1993). Alternatively, if transcription of the Izsp7O. 1 gene in murmne NIH3T3cells is abolished by the addition of an RNA-polymerase-II-specific inhibitor (5,6-dichloro-1 -Dt Protein kosmotropes (cf. chaotropes) are solutes which promote the proper folding of proteins.34ribofuranosylbenzimidazole, or DRB) to the culture medium during heat shock, then unregulated, runawaytranscription of the multicopy B2 heat-shock gene (which is transcribed by RNA polymerase III and thereforeDRB-insensitive) is observed (Price & Calderwood, 1992).The predicted coiled-coil domain of HSP7O (between amino acids 496 to 542) has been proposed to actas a site for the interaction of HSP7O and HSF (Hightower, 1991; Lupas et al., 1991). HSP7O is required for theimport of proteins into the nucleus, which implies that HSF-HSP7O complex formation at this stage is obligatory(Shi & Thomas, 1992). In addition, depletion of intracellular ATP levels is known to induce the heat-shockresponse, presumably by interfering with the recycling of HSP7O bound to cellular proteins other than HSF(Baler et al., 1992). This would decrease the ratio of free HSP7O to HSF in the cell and lead to an increase in theavailability of free HSF. Further studies will undoubtedly reveal more insights into the autoregulation of theheat-shock response (Craig & Gross, 1991; Yost et al., 1990a).C THE NEMATODE CAENORHABDITIS ELEGANS.1. Salient attributes.C. elegans is a nonparasitic, microbivorous soil nematode that has seven important attributes for amodel biological system: a small body size (.- 1 mm long), a short generation time (-. 3 days), a large brood size(-.300 eggs), tractable genetics (adults are self-fertilizing hermaphrodites that can give rise to males at lowfrequency), ease of propagation under laboratory conditions, limited complexity for a metazoan organism(-.1,000 somatic cells at maturity), and a transparent body (Brenner, 1974; Suiston & Brenner, 1974). C. eleganshas been studied extensively by researchers interested in behaviour, development, genetics, molecular biology,and neurophysiology (Kenyon, 1988; Wood, 1988).Two drawbacks to the use of C. elegans as an experimental system are a relative paucity of observablemutant phenotypes (e.g., compared to the fruit fly Drosophila melanogaster), and the difficulties that thecollagenous cuticle (surrounding the adult) and the chitinous shell (surrounding the egg) pose to biochemicalstudies that require subcellular fractionation (Wood, 1988). The former difficulty is less of a problem than it35used to be, due to the increasing research interest in C. elegans and the concomitant development of moresophisticated genetic techniques for maintaining the viability of mutant phenotypes (Wood, 1988).2. Studies of DNA-binding proteins and chromatin structure.Noteworthy achievements in this area of research include the purification and analysis of RNApolymerase II (Sanford et al., 1983, 1985), some preliminary work on the isolation of RNA polymerase IIregulatory factors (Okai, 1986a, 1986b), and the purification and analysis of C. elegans histones and nonhistonechromosomal proteins (Meheus & Vanfleteren, 1985; Vanfleteren, 1982; Vanfleteren & Van Beeumen, 1983;Vanfleteren et al., 1979). Changes in the overall structure of chromatin during development and changes inlocus-specific chromatin structure during heat-shock gene activation have also been studied (Dixon et al., 1990;Meheus & Vanfleteren, 1986).An in vitro transcription system for genes transcribed by RNA polymerase III has been developed usingcell-free extracts from C. elegans (Honda et al., 1986). Unfortunately there is no homologous in vitrotranscription system available for the study of C. elegans genes transcribed by RNA polymerase II.3. Isolation of genes that encode sequence-specific DNA-binding proteins.Although no sequence-specific DNA-binding proteins have been purified from C. elegans, severalgenes have been isolated that are believed to encode such proteins. Alternate splicing of mRNA derived fromthe sex-determining tra-1 gene produces two transcripts that encode zinc-finger proteins (Zarkower & Hodgkin,1992, 1993). Alternate splicing of mRNA derived from the egl-43 gene also produces two transcripts thatencode zinc-finger proteins (Garriga et al., 1993). Egl-43 proteins act as regulators of the migration ofhermaphrodite-specific neurons and of phasmid neuron development, and they are structurally similar to themurine Evi-1 proto-oncoprotein (Garriga et al., 1993). The sdc-1 gene also encodes a zinc-finger protein, andthis transcription factor is believed to regulate sex determination and dosage compensation in C. elegans (Nonet& Meyer, 1991).The product of the heterochronic gene lin-14 is a nuclear protein that acts as a developmental switch,and heterochronic mutations in lin-14 cause cells at a particular stage of development to adopt a fate that is36usually encountered either earlier or later in development (Ruvkun & Giusto, 1989). The product of the tin-31gene specifies three alternative cell fates in vulval development, and this protein belongs to the FINF-3lfork headfamily of transcription factors (Miller et al., 1993). The ett-1 gene, which is expressed in C. elegans embryos,encodes a protein which is similar in amino acid sequence to members of the (vertebrate) GATA transcription-factor family (Spieth et at., 1991). The Elt-1 protein is required for the expression of vitellogenin, tra-2, andmsp genes in C. elegans (Spieth et at., 1991).A large number of homeobox-containing genes have been cloned from C. elegans. The expressionproduct of the mec-3 gene contains a homeodomain, and is involved in the differentiation of touch receptorneurons (Way & Chalfie, 1988; Xue et at., 1992). Another homeobox-containing gene involved in thedevelopment of the nervous system is unc-4 (Miller et at., 1992). The C. elegans cell lineage and differentiationgene unc-86 is one of several developmental genes whose sequence similarity led to the identification of thehomeobox-containing “POU” domain (it-1, ct-1, LLnc-86; Finney et at., 1988; Herr et at., 1988; Xue et al.,1992). Posterior pattern fonnation and the migratory behaviour of neuroblasts in C. elegans are regulated by theposition-specific expression of the mab-5 gene (Costa et at., 1988; Salser & Kenyon, 1992). The lin-39 gene isrequired for midbody region-specific development, and both lin-39 and mab-5 contain Antennapedia-likehomeodomains (Clark et at., 1993; Salser & Kenyon, 1992). Other homeobox-containing genes believed to beinvolved in pattern formation include en-3, en-5, hom-2, hom-3, prd-i, and prd-2 (Kamb et al., 1989). TheC. elegans cell-lineage gene tin-il also encodes a homeodomain (Freyd et at., 1990). In addition to ahomeodomain, the mec-3 and tin-li genes encode a novel, cysteine-rich protein sequence which is believed tofunction as a metal-binding domain (Freyd et al., 1990). Other homeobox-containing genes isolated fromC. etegans include 15 genes designated ceh-i to ceh-15, and the ceh-19 gene (Burglin et al., 1989; Hawkins &McGhee, 1990; Naito et at., 1992; Schaller et at., 1990).4. Heat-shock-inducible gene expression.C. elegans induces the synthesis of heat-shock proteins when exposed to temperatures 10-15 degreesabove the normal growth temperature of 20 °C (Snutch & Baillie, 1983). The induction of the heat-shockresponse is rapid, and the expression of heat-shock genes is under both transcriptional and translational control37(Snutch & Baillie, 1983). All of the various classes of heat-shock proteins can be induced at all stages ofnematode development (Snutch & Baillie, 1983). The most abundant heat-shock proteins are the expressionproducts of members of the hsp7O gene family (Snutch & Baillie, 1983; Snutch et aL, 1988).The Jispl6 gene family in C. elegans consists of two identical pairs of divergently transcribed geneslocated at one chromosomal locus, and a third pair of divergently transcribed genes located at a differentchromosomal locus (Candido et al., 1989). These two hspl6 gene pairs are expressed at very different levelsduring heat shock, despite the highly conserved sequences of the genes and their promoters (Jones et al., 1989).The differential regulation of these genes is believed to reflect differences in locus-specific chromatin structureduring heat shock. (Dixon et al., 1990).The C. elegans genome contains a moderately repetitive DNA sequence family with a consensussequence of a9 5 A’ 00 5 T5 T50T901 1 00 c90T5 A5 0, where the statistically observednucleotide frequencies (as percentages) are indicated by superscripts (La Volpe et a!., 1988). Multiple copies ofthis 10-bp repeating unit are found at various locations dispersed throughout the C. elegans genome (La Volpeet al., 1988; Naclerio et a!., 1992). Although these repetitive, HSE-consensus-like sequences (now designated asmembers of the RcC9 family) do not appear to direct the heat-inducible transcription of flanking DNA inC. elegans, a 242-bp copy of this repetitive DNA element did confer heat inducibility upon a reporter gene whenintroduced into yeast cells (La Volpe et a!., 1988; Naclerio et a!., 1992). The existence of a moderatelyrepetitive DNA family such as RcC9 is without precedent in eukaryotic genomes. The effect of these HSEislands on the transcriptional regulation of the heat-shock response in C. elegans is unknown.D. SUMMARY OF THESIS OBJECTIVES FOR PART 1.The focus of this work is on the detection and purification of sequence-specific DNA-binding proteins,and in particular, the study of a putative ceHSBF activity in the nematode C. elegans. No sequence-specificDNA-binding proteins have been isolated from C. elegans and studied at the biochemical level, despite theinterest that this species holds for developmental biologists, geneticists, molecular biologists, andneurophysiologists. One reason for this situation is a lack of reliable experimental methods to perform such38studies. Most of the studies of sequence-specific DNA-binding proteins in metazoan organisms have usedtissue-culture cells as a biological starting material. There are no such cell lines available from C. elegans.Furthermore, the procedures developed for the subceflular fractionation of tissue-culture cells differ substantiallyfrom the procedures appropriate for C. elegans. I hope that the experimental procedures described herein willfacilitate the study of sequence-specific DNA-binding proteins in C. elegans.As mentioned at the beginning of this introduction, there are compelling reasons for the intenseresearch interest in the study of eukaryotic, sequence-specific DNA-binding proteins. Unfortunately, thedevelopment of improved methods for the detection and purification of members of this important class ofproteins has not really received commensurate attention. In addition, some of the EMSA procedures and resultsreported in the literature are of potential concern. For example, one potential concern is the use of extremelylow ionic-strength buffers during electrophoresis (with I < 0.015 M; e.g., Fried & Crothers, 1981; Strauss &Varshavsky, 1984). Thus although this work focuses on the study of a putative sequence-specific DNA-bindingactivity from C. elegans, I hope that some of the experimental approaches described herein will assist similarstudies in other biological systems.39CHAPTER 1.2. MATERIALS AND METHODSA. MATERIALS USED IN THIS THESIS (PARTS 1 AND 2).ACS reagent grade chemicals and Difco powdered nutrients were used to prepare culture media.Analytical reagent grade chemicals and organic solvents were used for most other purposes, and these productswere used without further purification. Some noteworthy suppliers of these and other reagents are mentionedbelow. Unless specified otherwise, acids and bases were obtained in their un-ionized forms.Aldrich supplied TCBA. BDH Canada Ltd. supplied 6% sodium hypochlorite solution. BethesdaResearch Laboratories supplied “ultrapure” grade formamide and redistilled phenol. Redistilled phenol wasliquefied and then equilibrated with TRIS-HC1 pH 8.0 as described (Sambrook et at., 1989). Fisher Scientificsupplied methanol (HPLC grade), triethylamine (HPLC grade), and pH-meter standardization buffers. Flukasupplied the following reagents, which were generally the highest purity available: acrylamide; 4-aminobutanoicacid; biological buffers; cyclam; DABCO; EGTA; Et4NBr; the monopotassium salt of L-glutamic acid(KHG1u.H20);2-mercaptoethanol; N, N’-methylene-bis-acrylamide; phosphate salts; piperazine; and sucrose.Sigma supplied antifoam A emulsion, BSA (1.0 mg/mL standard for protein assays), DAPI, dimethylsulfoxide (tissue-culture grade, ampouled in 5-mL aliquots under nitrogen), “ultrapure” DNA (from calf thymus,C. peifringens and M. luteus; 42, 26, and 72% G+C; Cat no. D4764, D5139, and D5014, respectively), DTT,electrophoresis buffer ingredients (except KHG1u.H20), heparin (sodium salt, grade I-A, 170 USP units/mg)Hoescht 33258 dye, n-octyl f3-D-glucoside, PEG 8000 (DNase-free, molecular biology grade),polyacrythydrazido agarose, protease inhibitors, and yeast tRNA.Boehringer Mannheim supplied BSA (DNase-free, molecular biology grade), proteinase K, somerestriction endonucleases (along with recommended buffers), and Triton X-100 (specially purified for membraneresearch, ampouled in 5-mL aliquots under nitrogen). Bio-Rad supplied ammonium persulfate, all of thechromatography resins, a Bradford protein-assay kit, Poly-Prep chromatography columns, protein molecularweight standards for electrophoresis (silver-stain high-Me markers), a protein (PAGE gel) Silver-Stain Plus kit,40SDS-PAGE buffer concentrate, and TEMED. Millipore supplied disposable 0.4- and 2.O-mL ultrafiltration units(Ultrafree MC and CL, low protein-binding cellulose, 10,000 nominal Mr limit).Amersham Hyperfilm MP X-ray film was used for autoradiography. Fuji Neopan film (ASA 1600)was used for microphotography, and a standard stage micrometer (0.10 and 0.010 mm divisions) was used forthe calibration of microphotographs. New England Nuclear supplied Cronex Lightning Plus intensifyingscreens, NENSORB-20 nucleic acid purification columns, and radioisotopes. Polaroid Type 57 film was usedfor the photography of electrophoresis gels.Pharmacia supplied ATP and dNTPs (obtained as neutralized 100 mM stock solutions),poly(dI-dC).poly(dI-dC) (lots BG7880204 and CJ7880107; average fragment sizes of 8,600 and 1,200 bp; A260units/mg values of 10.0 and 12.7; designated herein as lots I-i and 1-2, respectively), pdN5, pdN6 andribonuclease A. Stock solutions of ribonuclease A were heated to remove any possible traces of DNase activity(Sambrook et al., 1989). New England Biolabs supplied most of the restriction endonucleases (along with theirrecommended buffers). New England Biolabs and Oligos Etc. Inc. (Wilsonville, Oregon, U.S.A.) synthesizedall of the oligonucleotides used in this thesis. United States Biochemical Corporation supplied Sequenaseversion 2.0.Nematodes grown in liquid culture were fed E. coli cells (“washed”, 3/4 log phase, strain MRE 600)that were obtained as a frozen paste from the Grain Processing Corporation (Muscataine, Iowa). This materialshould never be used for this purpose because this material consistently gives rise to very seriouscontamination problems. Uncontaminated E. coli cells (prepared by in-house fermentation as recently aspossible prior to use) are essential for the proper growth of C. elegans in large-scale liquid cultures (Gbewonyoet al., 1994; Suiston & Hodgkin, 1988).B. ROUTINE EXPERIMENTAL PROCEDURES USED IN THIS THESIS (PARTS 1 AND 2).1. Preparation of solutions.Distilled water was used in culture media and in solutions used to prepare eggs. Type I water, preparedby a Milli-Q (Millipore) or Nanopure (Barnstead) apparatus, was used in all other solutions.41Table 2. Hydrogen-ion buffered solutions formulated for this work.Name Solute Concentration pH CommentsESB KC1 100 mM 7.5 a.Egg storage MgCl 5.0 mMbuffer HEPES 10 mMNaOH to adjust pHBuffer X KC1 75 mM 7.5 a, c.MgC1 2.5 mMEGTA 1.0 mMHEPES 20 mMNaOH to adjust pHBuffer X.40 Buffer X 7.5 b, d, e, f.Preparedjust before OG 0.40 % wlvuse from bufferX DTr 2.0 mME-64, Pepstatin A 20 jiM eachPMSF 0.4 mMBuffer Y KC1 75 mM 7.5 a, c.MgCl 2.5 mMEGTA 0.5 mMHEPES 10 mMNaOH to adjust pHBuffer Y.02 Buffer Y 7.5 b, d, e, f.Preparedjust before OG 0.02 % wlvuse from buffer Y DTF 1.0 mME-64,PepstatinA 10 jiMeachPMSF 0.2 mMBuffer Z KC1 900 mM 7.5 a, c.HEPES 10 mMNaOH to adjust pHBuffer A KC1 120 mM 7.1 a, b, c.MgCl 3.5 mMEGTA 1.0 mMBIS-TRIS 20 mMHEPES 20 mMBuffer A.02 Buffer A 7.1 b, d, e, f.Preparedjust before OG 0.02 % wlvuse from bufferA DTl 1.0 mMProtease inhibitors 10 jiMBuffer A.42 Buffer A.02 7.1 b, d, e, g.Preparedjust before OG 0.42 % wlvuse from bufferA.02 DTT 11 mMBufferfi KC1 1.5 M 7.1 a,b.Na2EDTA.2H20 30 mMBIS-TRIS 60 mM42Name Solute Concentration pH CommentsKBHE KC1 100 mM 7.0 a, b.BIS-TRIS, HEPES 20 mM eachNa2EDTA.2H20 1 mMBHE BIS-TRIS, HEPES 20 mM each 7.0 a, b.Na2EDTA.2H20 1 mMKGBH (8x) KHGIu.H20 800 mM ..7.3 (8x) a, b, c.BIS-TRIS, HEPES 160 mM each -7.0 (lx)KGBM (8x) KHG1u.H20 1.2 M -6.6 (8x) a, b, c.BIS-TRIS, MES 400 mM each -.6.3 (lx)KGTH (8x) KHG1u.H20 1.2 M -.8.3 (8x) a, b, c.TRIS, HEPES 400 mM each —7.9 (lx)CWB KC1 150 mM 7.0 a, b, d.Column BIS-TRIS, HEPES 20 mM eachwash buffer Na2EDTA.2H20 1 mMOG 0.20%w/vCEB-2000 ICC! 2.0 M 7.0 a, b, d, h.Column BIS-TRIS, HEPES 20 mM eachelution buffer Na2EDTA.2H20 1 mM(see note “it” below) OG 0.20 % wlvSB NaH2PO4.2H20 -.120 mM 7.0 a, b, I.Stop buffer Na2HPO4 -.380 mMTP (1.25x) KCI 60 mM 4.20 a, j, k.TCBA 10 mMPEG 8000 15 % wlvPiperazine to adjust pHTC (1.25x) TP with cyclam replacing piperazine. 4.20 a, j, k.(a) Filter sterilize.(b) Should not need to adjust the pH (n.b. which components are salts and which are un-ionized).(c) Store in the dark at4°C.(d) Add OG from a freshly prepared 20% wlv stock solution just before use.(e) Add DTF from a freshly thawed aliquot of a 1.0 M stock solution just before use.(f) Thaw aliquots of stock solutions of protease inhibitors and add just before use.(g) OG and Dfl’ concentrations shown include conthbutions from buffer A.02.(h) CEB-3, CEB-, CEB-j, and CEB-2 differ only in the concentration of KC1 (given in)(i) Contains 0.50 M phosphate and 0.88 M sodium ions.(j) Store in the dark at 20-22 °C and use within 24 h after preparation.(k) Prepamd at 1.25x the final concentration in the immobilization reaction.43Beckman (model 3500) and Radiometer (model PHM 82) pH meters equipped with a Corningsemimicro combination electrode (model no. 476541) were used for pH measurements, and the pHmeter/electrode combination was standardized prior to use. Any buffered solution that contained PEG had thissolute added before the pH was adjusted. All pH measurements were made at 20-22 °C, regardless of thetemperature at which the buffer was to be used. Coarse pH adjustment (to within 0.04 to 0.2 pH units,depending on the solution) was carried out with solutions at about 90% of their final volumes. Final pHadjustment (to within 0.01 pH units) was made with solutions at their final volumes, as the amount of acid orbase added at this stage was negligible. These details are important to note, as several of the buffer solutes thatwere used have significant dpKa/dT values; furthermore, PEG has a pronounced, concentration-dependent effecton the pH of buffered solutions (Atha & Ingham, 1981; Eichele et al., 1978).2. Liquid handling and plastic ware.Gilson adjustable, automatic micropipettors were used for routine liquid handlingt. An adjustable“Ultramicropipettor” (Eppendort) was used for handling volumes from 10.0 to 0.5 microlitres, and this devicetwas used for all EMSAs.Unless specified otherwise, the following designations pertain to disposable or durable plastic ware(which was either radiation-sterilized by the manufacturer or autoclaved): “microcentrifuge tube” (1 .5-mLpolypropylene tube with attached snap-cap), “cryotube” (1 .8-mL nominal volume, round-bottomed, allpolypropylene cryogenic storage vial with externally threaded screw cap, Nunc Cat no. 375418), “14-mL tube”(round-bottomed, polypropylene clinical centrifuge tube with polyethylene snap cap, Falcon Cat no. 2059),“40-mL Oakridge tube” (round-bottomed, polyallomer centrifuge tube with polypropylene screw cap), “50-mLtube” (conical-bottomed, polypropylene clinical centrifuge tube with polyethylene screw cap), “250-mL bottle”(flat-bottomed, polyallomer centrifuge bottle with polypropylene screw cap).t The instruments used were recently purchased. They were calibrated to deliver the volumes reported, and toan accuracy consistent with the number of significant figures reported. The accuracy and precision of theseinstruments vary depending on the volume dispensed; the temperature, density, and viscosity of the solution; andthe type of tip used (and how properly that tip was affixed). The documentation supplied by each manufacturerprovides a full discussion of the accuracy and precision of these instruments.443. Aseptic technique.Sterile solutions were used in all cases, except when harvested nematodes were washed prior to alkalinehypochiorite treatment. Most of the solutions were sterilized using disposable sterile-filtration devices (whichhad nominal pore sizes of 220 nm unless specified otherwise) obtained from several different vendors.Supernates were routinely aspirated using aseptic technique. This involved the use of nonsterile, glassPasteur pipettes and sterile, 200-.tL (regular) or 10-.tL (Ultramicropipettor gel-loading) size, disposablepolypropylene pipette tips. The disposable tips were retained on the tip of the Pasteur pipette by the vacuumsource used for aspiration.A tissue-culture facility laminar-flow hood provided a sterile environment for all critical manipulationsand experiments, including those involving the handling of chromatography resins.4. Mixing by rotation.The term “mixed by rotation” denotes “parallel” rotation (tube axis parallel to a nearly horizontal axisof rotation) at 10 rpm using a mixing wheel (Roto-Torque, Cole-Parmer).5. Small-scale degassing of solutions or chromatography resins.The term “the solution (or resin) was degassed” denotes an aseptic procedure where the material inquestion (up to 12 mL) was carefully deposited at the bottom of a 50-mL tube and then degassed for exactly5 mm using a water aspirator as a vacuum source. A clean, oversized, single-holed rubber stopper (pierced witha 450 angle-shaped piece of bent glass tubing that was connected to the vacuum line) was used to seal theopening of the 50-mL tube. The tube was agitated for the first two minutes of degassing to minimize bumping.WARNING: The 50-mL tubes used in this procedure are not rated for use under reduced pressure. Although Ihave never had any problems using this technique, caution and the use of adequate eye protection are essential.6. Microcentrifugation.The term “microcentrifuged” denotes centrifugation in an Eppendorf microcentrifuge at 12,000 g.457. Miscellaneous procedures for handling DNA.a. DNA precipitation and resuspension.Standard conditions for the precipitation of DNA were used, i.e., the addition of 0.1 vol of 3.0 MNaOAc pH 5.2 and either 2 vol of EtOH or 1 vol of 2-propanol. After an appropriate time interval, precipitatedDNA samples were centrifuged at 12,000 g for 5 to 60 mm. High-Mr DNA (>200 bp) was precipitated andcentrifuged at either 4 °C or at 20-22 °C (centrifuged at 20-22 °C if the DNA was precipitated from solutionscontaining telraalkylammonium salts), and then all traces of the supemate were removed. High-Mr DNA pelletswere rinsed with 70% EtOH, pelleted 2 to 30 mm, and then all traces of the supernate were removed after eachrinse. These pellets were dried for 10 mm in a clean desiccator (i. e., containing no desiccant) with a wateraspirator as a vacuum source. DNA pellets were resuspended with TE (at 20-22 °C) or 10 mM KC1 (on ice), andthe resulting solutions were stored at -20 °C.Poly(dI-dC).poly(dI-dC) used in EMSAs was resuspended with KBHE, left for several days at 4 °C,quantitated spectrophotometrically, and then adjusted to a concentration of 2.0 mg/mL with KBI{E.Special precautions were taken when handling the HSE probe. This DNA was precipitated at least16 hours at 4 °C, microcenthfugecl for 30 to 60 mm at 4 °C (or at 20-22 °C if the DNA was precipitated fromsolutions containing tetraalkylammonium salts), and then all traces of the supernate were removed. The pelletedprobe DNA was rinsed with 99% EtOH (or 70% EtOH if the DNA was to be resuspended with 10 mM KC1),microcentrifuged for 10 to 30 mm, and then all traces of the supemate were removed after each rinse. The DNApellet was air-dried for 30 mm in a sterile environment, resuspended on ice with 10 mM KC1 or TE pH 7.4, andthe resulting solutions were stored at -20 °C or 4 °C as specified. The precaution of briefly air-drying the probewas warranted by a report of spurious EMSA results obtained using excessively dried, low-Mr DNA probes(Svaren et a!., 1987).Random-sequence oligonucleotides (pdN5 or PdN6) were obtained as lyophilized powders, and wereresuspended to 2.0 mg/mL in BHE buffer.46b. DNA assays.DNA was assayed by UV-ABS spectrophotometry on a dual-beam, Shimadzu UV 160 instrument.UV-ABS DNA assays were performed using an appropriate blank, and these assays involved running a spectrumfrom 3 20-250 urn, and printing out the absorbance value at each 10-nm interval. Unless specified otherwise(e.g., for poly(dI-dC)•poly(dI-dC) or the duplex, 5 1-bp HSE probe), 1.00 A260 unit (the amount of light-absorbing solute in a 1.00 mL solution that yields an A260 of 1.00 in a 1.00-cm path length) was assumed toequal 50 micrograms of double-stranded DNA (Sambrook et al., 1989).Spectrofluorometric assays using Hoescht 33258 dye were performed on an SLM/Aminco SPF-500Cinstrument (Labarca & Paigen, 1980). The spectrofluorometer excitation and emission maxima (and associatedbandpass values) were set at 356 (20 urn) and 458 nrn (7.5 nm), respectively. Fluorescence enhancement wasmeasured by recording the sample/reference ratio (relative fluorescence) of the assay buffer before and afteraddition of the DNA solution.C PREPARATION OF NUCLEAR EXTRACTS FROM C. ELEGANS EGGS.1. Large-scale liquid culture of C. elegans.The N2 strain of Caenorhabditis elegans var. Bristol was used for all experiments. SynchronousC. elegans cultures (0.25 to 3.0 liires)t were grown at 20-22 °C with vigorous aeration in supplemented Basal Smedium (Brenner, 1974). Each culture was started with 1 g of (floated) eggs per litre, together with 7 to 10%w/v E. coli and adequate amounts of antifoam A emulsion. After 3 to 4 days, gravid adult nematodes wereallowed to settle at 4 °C prior to extensive washing with cold 0.14 M NaC1. Nematode eggs were prepared fromgravicl adults essentially as described (Emmons et al., 1979; see below for a large-scale version of thisprocedure). These eggs were used to start another culture or were subjected to heat-shock (or control)incubatious as described below.t The culture vessels, which I had to design and build myself, are not described here. Proper fermentationequipment is readily available, and under no circumstances should nematodes be grown in large-scale liquidculture using improvised equipment. Bioreactor-based cultivation and downstrearn processing of large-scalecultures of C. elegans are described in Gbewonyo et al. (1994).472. Large-scale preparation of C. elegans eggs and exposure to heat shock.Washed, gravid nematodes were floated in cold 30% wlv sucrose, washed again with cold 0.14 MNaC1, and then distributed in 30- to 40-g aliquots in 250-mL bottles. Cold, alkaline hypochlorite solution(prepared the day of the experiment and prechilled on ice) was added (200 mLlbottle), and the bottles wereshaken for 4 mm at 20-22 °C. Eggs and undisrupted adult bodies were pelleted for 2.5 mm at 3,000 rpm using aBeclcman JA 14 rotor and 12-2 1 centrifuge at 20-22 °C. During the hypochionte disruption procedure all large-volume centrifugation runs were performed using this rotor-centrifuge combination, and the brake was set at themaximum setting. The supernate was carefully poured off and replaced with cold alkaline hypochlorite solution(200 mL/bottle). The bottles were shaken until disruption of the adult bodies was complete (after 1-2 mm;disruption must be monitored by microscopy). The eggs were pelleted as before, except that this time (and aftersubsequent washes) the supemate was carefully removed by aspiration.Eggs were washed three times with cold 0.14 M NaCl, and then floated in cold 30% w/v sucrose.Floated eggs were diluted and washed with cold 0.14 M NaCl, and the wet weight of the pelleted eggs wasdetermined. Approximately 15 g of clean, viable eggs was obtained from each 3-litre culture. The eggs wereresuspended in a small amount (<50 mL) of egg storage buffer (ESB), a fraction was retained to start a successorculture, and the remainder (30-50 mL) was subjected to control (20 °C) or heat-shock (33 °C) incubations (seebelow). At this point a very small amount of the residual egg suspension in the bottle was used for a“preshock” viability test (i.e., about 300 to 500 eggs were spotted on a plate).The egg suspension (30-50 mL in ESB) was added to 200 inL of ESB (pre-incubated at either 20 or33 °C) in a 2-litre Erlenmeyer flask. The flask was then shaken in a 20 or 33 °C water bath for 20 mm at 250rpm. After the 20-mm incubation was over, the eggs were collected in five 50-mL tubes, pelleted at 20-22 °Cfor 3 mm at 2,000 rpm in a clinical centrifuge, and the supernate was removed. ESB was added to give a 1:1 (wteggs:vol ESB) suspension, and then the well-mixed suspension was dripped into a plastic strainer submerged inliquid nitrogen. The quick-frozen egg pellets were collected (by simply removing the strainer from the liquidnitrogen), transferred to labeled containers, and then stored at -70 °C. At this point a “postshock” egg viabilitytest was carried out (using the residual egg:ESB suspension) as described above.48Buffer A(l8OmL)120 mMKCI1 mM EGTA3.5 mMMgCI220mM Bis-Tris20mM HepespH?.lAdd 0.001 vol of:I1.OOMDTr20% wlv OG10mM protease inhibitorsliiiBuffer A.02—Buffer A.02:(7.0 mL)Add 0.01 vol of1.OOMDTTAdd 0.02 vol of20% w/v OGBuffer A.42:as per A.02,but with0.42% wlv OG11 mMDTTri I I I I‘J Add 4 g of 50% w/v Remove a 0.4% aliquotfrozen eggs of egg suspensionThaw eggs by mixing Pellet eggs 4 nun at 2,000 rpm;aspirate the supernate46 mL ADisrupt cellsIIIIII_______(100 strokes)I I Dilute the homogenate_Ijlj(7OmLofA.02)Pellet debris4 mm at 2,000 rpmTransfer the supennateto two Oaknidge tubesPellet nuclei 8 mm at 4,000 rpm;aspirate the supernateResuspend the pelletsthen wash the nuclei(39 mL of A.02 per tube)Pellet nuclei 8 miii at 4,000 rpm;aspirate the supernateResuspend nuclei with A.02to a total of 1.00 mLRemove a 1.2% aliquot ofresuspended nucleiAdd 0.2 vol of Buffer B:1.50 MKCI 60mM Bis-Tris3OmMEDTA pH7.1Extract nuclei 30 miii with stimug;microcentrifuge debris 5 mmFilter, aliquot, and freeze the supemate;store at -70°CResuspend eggs(7.0 mL of A.42)Transfertostainlesssteel 111111Dounce homogenizerFigure 2. The A-protocol for the preparation of nuclei and nuclear extracts from C. elegans eggs.49The viability of the eggs (pre- or post-shock) was essentially quantitative (>98%), and the putativeceHSBFm activity proved to be very stable to prolonged storage at -70 °C. The EMSA experiments wereperfonned using material that had been stored at -70 °C for up to 3 years.3. Preparation of 0.35 M KC1 nuclear extracts: The A-protocol.This procedure is summarized in Fig. 2 and described in detail below.Stock solutions (l000x) of protease inhibitors (10 mM of the active inhibitor; some were not suppliedas pure compounds) were prepared in either water (E-64), methanol (bestatin), or dimethyl sulfoxide (leupeptinor pepstatin A). With the exception of bestatin, the protease inhibitors chosen were selected based on the resultsof Sarkis et al. (1988). These stock solutions were aliquoted, stored at -70 °C, and aliquots were thawed justbefore use. Buffer A.02 was prepared from buffer A just before use by supplementing 180 mL of buffer A in apolypropylene beaker with 0.001 vol of the following stock solutions: 1.0 M DTr; 20% w/v 06; and 10 mMsolutions of bestatin, E-64, leupeptin, and pepstatin A. Buffer A.42 was prepared by pouring 7 mL of freshlyprepared buffer A.02 into a 14-mL tube, then adding 0.01 vol of 1.0 M DTI’ and 0.02 vol of 20% w/v OG andgently mixing the solution. All buffers were prechilled and experimental manipulations were performed onice or at 4 °C unless specified otherwise.Four grans of the frozen, 1:1 w:v egg:ESB mixture (i.e., -2 g of eggs in a total of 4 mL of suspensionwhen thawed) was added to a 50-mL tube that contained 46 mL of unsupplemented buffer A. The eggs werethawed rapidly by mixing, and then 0.200 mL of the suspension was quantitatively transferred (i.e., washingswere added after rinsing the pipette tip three times with unsupplemented buffer A) to a microcentrifuge tube.The eggs in this 0.4% aliquot were microcentrifuged (1 mm), the supemate was carefully removed, and the eggsample was stored at -20 °C.The following steps of the protocol involve differential centrifugation. To prevent disruption ofpelleted fractions, no braking of the IEC or the Sorvall RC-5 centrifuge was employed. Centrifugation runsperformed with an IEC benchtop centrifuge were (at time t=0) accelerated to 2,000 rpm as rapidly as possible(i.e., the speed setting was immediately increased to the maximum value), adjusted to 2,000 rpm (i.e., byreducing the speed setting once 2,000 rpm was reached), and then (at time t=4 mm) the speed setting was50immediately reduced to zero. At a setting of 2,000 rpm the nominal and the measured speed (detennined using astroboscope) of the JEC centrifuge agreed to within 50 rpm.The thawed eggs in the 50-mL tube were pelleted 4 mm at 2,000 rpm in an IEC HN-S II modelbenchtop centrifuge and the supernate was removed by aspiration. The 7-mL aliquot of buffer A.42 was used toresuspend and transfer the eggs to a 15-mL stainless steel Dounce homogenizer (Wheaton). The eggs werehomogenized using 100 strokes. A straight, vertical motion was used on the downward half of each stroke. Thepestle was pulled back at a slight angle (with slight side-to-side rotation) on the upward half of each stroke. Thehomogenate and washings from the homogenizer were transferred to a 250-mL polypropylene beaker using 70mL of buffer A.02, mixed well, and then transferred to two 50-mL tubes.Debris was pelleted in an IEC HN-S II centhfuge (4 mm at 2,000 rpm, with the tube caps removed).The supernate was very carefully decanted into two 40-mL Oakridge tubes. Nuclei present in the transferredsupernate were then pelleted using a Sorvall HB-4 swinging-bucket rotor and RC-5 centrifuge (8 mm at 4,000rpm). The supernate was carefully aspirated, each pellet was resuspended using a micropipettor and —0.8 mL ofbuffer A.02 (—40 pipettings per pellet; avoid introducing air into the nuclei suspension), and then 39 mL ofbuffer A.02 was added to each tube. Resuspended nuclei were washed by mixing, pelleted, and the supernatewas removed as before. Each pellet was carefully resuspended using a micropipettor and 0.3 mL of buffer A.02,and then the nuclei were pooled in one tube.The remaining experimental procedures were carried out at 20-22 °C. The volume was measuredwhile transferring the nuclei suspension to a 14-mL tube containing a 3x10 mm “flea” stir bar. The volume ofthe suspension was adjusted to 1.0 mL with buffer A.02. The tube was clamped about 1 cm above a magneticstir plate, and the nuclei were stirred moderately (at 0.6x max. speed). A 12-iL aliquot of nuclei was transferredto a microcentrifuge tube and stored at -20 °C. An 8-.iL aliquot was also removed, mixed with 0.5 jiL of DAPI(1 mg/mL in water), stored on ice, and later used to examine the nuclei by DIC and epi-fluorescence microscopyon a Zeiss IM 35 inverted microscope. A 0.2-vol aliquot of 20-22 °C buffer B was then added dropwise, withstirring, to the nuclei. Moderate stirring (at 0.6x max. speed) was continued for 30 mm thereafter. The crudenuclear extract was transferred to a microcentrifuge tube, and debris was microcentrifuged for 5 mm. The51supernate was transferred to a 1O-mL disposable polypropylene syringe equipped with a filtration device(450-nm pore size, low protein-binding, 13-mm dia., Millex GV13,Millipore). The extract was filtered, mixedgently, divided into 50-pL aliquots in 0.5-mL microcenthfuge tubes (or kept in one single -.1 .0-mi. aliquot in a1.5-mi. microcentrifuge tube for the affinity chromatography experiments), frozen quickly in a dry ice/ethanolbath, and then stored at -70 °C.Although the mortar and the tip of the pestle of the Dounce homogenizer are made of stainless steel, theshaft of the pestle will corrode after exposure to moisture; it is therefore recommended that this instrument bewashed and dried as soon as possible after use.4. Preparation of 0.35 M KCI nuclear extracts: The X-protocol and the AP-protocol..The differences between the X-protocol and the A-protocol are noted below.Stock solutions of (10 mM) of E-64 and pepstatin A, as well as PMSF (200 mM in 2-propanol), wereprepared and stored as described earlier. Buffer X.42 was prepared from buffer X just before use bysupplementing 7 mL of buffer X in a 14-mL tube with 0.002 vol of 1.0 M DTr, 10mM E-64, 10 mMpepstatin A, 200 mM PMSF and 0.02 vol of 20% wlv OG, and gently mixing the solution. Buffer Y.02 wasprepared from buffer Y just before use by supplementing 150 mL of buffer Y in a polypropylene beaker with0.001 vol of 1.0 M DTT, 20% w/v OG, 10 mM E-64, 10 mM pepstatin A, and 200mM PMSF. Stock PMSF wasadded dropwise to rapidly stirring solutions.The X-protocol was carried out using two grams of the frozen, 1:1 w:v egg:ESB mixture (i.e., .4 g ofeggs in a total of -2 mL of suspension when thawed). The eggs were thawed using 43 mL of unsupplementedbuffer X, pelleted, the supernate was removed, the eggs were resuspended and transferred to the stainless steelDounce homogenizer using 7 mL of buffer X.42, and the eggs were homogenized (60 strokes only) essentiallyas described in the A-protocol. The homogenate was diluted with 35 mL of buffer Y.02, debris was pelleted in a50-mL tube, the supernate was transferred to a single 40-mL Oakridge tube, and then the nuclei were pelletedand the supernate discarded as described earlier. The nuclei were resuspended and washed with buffer Y.02 asdescribed earlier, except that in the X-protocol the nuclei were washed twice. The nuclei were resuspended to atotal volume of 0.42 mL with buffer Y.02 and then set up for the extraction (i.e., with stirring) as described52earlier. A 15-l.tL aliquot of resuspended nuclei was removed for qualitative microscopic analysis; noquantitative analysis of nuclei yield was attempted. The following solutions were added dropwise to the stirringnuclei: 15 .iL of 0.10 MNa2EDTA, pH 7.5 and then 0.21 mL of buffer Z. The nuclei were stirred vigorously for60 mm at 20-22 °C, and then the nuclear extract was clarified, filtered, quick-frozen, and stored at -70 °C asdescribed in the A-protocol.The differences between the AP-protocol and the A-protocol are noted below.Stock solutions of (10 mM) of E-64 and pepstatin A, as well as PMSF (100 mM in 2-propanol), wereprepared and stored as described earlier. The fmal concentrations of DTT, E-64, pepstatm A, and PMSF in bothbuffer A.02 and “A.42” were 1 mM, 10 pM, 10 pM, and 100 jiM, respectively. The protease inhibitors bestatinand leupeptin were not used in the AP-protocol. Buffer A.02 and “A.42” contained 0.02 and 0.22 % w/v OG,respectively.The AP-protocol was carried out using two grams of the frozen, 1:1 w:v egg:ESB mixture (i.e., 1 g ofeggs in a total of —2 mL of suspension when thawed). The work up and washing of the nuclei in the AP protocolis the same as in the X-protocol, except that the A-protocol buffers were used. In the AP-protocol the purifiednuclei were resuspended to a final volume of 0.52 mL with buffer A.02, then extracted as in the A-protocol. Thenuclear extract was clarified, filtered, quick-frozen, and stored at -70 °C as described in the A-protocol.5. Quantitative analyses of A-protocol nuclei and nuclear extracts.The nominal concentration of KC1 in the nuclear extracts was assumed to be 350 millimolar. Theprotein concentration in the nuclear extracts was determined using BSA as a standard (Bradford, 1976).The yield of nuclei was estimated by a standard DNA assay (Labarca & Pamgen, 1980). The followingreagents were added to each (-.10-j.LL) egg pellet or aliquot of nuclei: 760 pL of water, 20 j.tL of 0.5 MNa2EDTA pH 8.0, 100 jiL of 0.2 M TRIS-HC1 pH 8.0, 10 pLL of 10% w/v SDS, and 100 j.tL of 10 mglmLDNase-free proteinase K. The reactions were gently but thoroughly resuspended, transferred to cryotubes, andthen mixed by rotation for 16 hours at 50 °C. It is very important to properly resuspend the reactions prior toincubation. Aliquots (10 pL, in duplicate, each of which contains about 200 ± 50 ng of DNA) from these53digestions were then assayed using 2.0 mL of high-salt assay buffer (2.0 M NaCl; 50 mM phosphate, sodiumsalt, pH 7.4; 1 jig/mL Hoescht dye 33258). Calf thymus DNA (50 .tg/mL in TE pH 7.4) was used as a standard.The apparent relative yield of nuclei was calculated by dividing the value of the standard DNA assayobtained using the (1.2%) nuclei sample by three times the value of the standard DNA assay obtained using the(0.4%) egg sample, and then expressing this fraction as a percentage. The apparent absolute yield of nuclei (asmg of DNA per gram of wet tissue) was calculated by dividing the value (as mg of DNA) of the standard DNAassay obtained using one percent of the (1.2%) nuclei sample by 2.4 x grams. The denominator, 2.4 x i0grams, is three times the mass of the eggs in one percent (i.e., 10 iL assayed Out of each 1 .0-mL proteinase Kdigest) of the 0.4% egg sample (after thawing 2.0 g of eggs in 50 mL, a 0.200-mi. aliquot was removed).D. ELECrROPHORETIC MOBILITY-SHIFTASSAYS (EMSAs).1. Oligonucleotides used to form HSE probes.Commercially obtained oligonucleotides (see Fig. 3) were resuspended to nominal concentrations of0.1 mM in water. Oligonucleotide solutions were filter-sterilized and their concentrations were determinedaccurately by UV-ABS spectrophotometry. The molar extinction coefficients of the oligonucleotides hse-A,hse-T, hse-R and hse-Y were calculated to be 5.72, 5.48, 5.69, and 5.40 x 10M4cm,respectively (Sambrooket al., 1989).The annealing reactions (0.45 mL) were set up in 0.5-mL microcentrifuge tubes and containedequimolar concentrations of complementary oligonucleotides in 2.4 M Et4NBr, 10 mM NaEPPS, 1 mMNa2EDTA, pH 8.0. The oligonucleotide dissociation temperature (Td) for the 5 1-bp HSE duplex (47 bp ofwhich are double-stranded) in 2.4 MEt4NBr was estimated to be 53 °C, i.e., 30 °C below the corresponding Tdin 3.0 M Me4NC1 (Jacobs et al., 1988; Orosz & Wetmur, 1977; Wood et al., 1985). The annealing reactiontemperature was held 4 mm at 80 °C, decreased to 48 °C at 2 °Clmin, held 2 hours at 48 °C, decreased to 28 °Cat 1 °C/min, held 4 hours at 28 °C, and then decreased to 12 °C at 1 °C/min using an Ericomp “thermal cycler”apparatus. The annealing reactions were then cooled to 4 °C on ice, and the contents were transferred to 1.5-mi.microcentrifuge tubes. Fifty microlitres of 3.0 M NaOAc pH 5.2 and 1.0 mL of EtOH were added and then the54DNA was precipitated overnight. After pelleting of the DNA and removal of all traces of the supernate, eachDNA pellet was rinsed three times with 1.0 mL of 70% EtOH. The dried DNA pellets were then resuspendedwith 10 mM KC1 to a nominal concentration of 2.0 mg/mL. UV-ABS spectra were taken, and then the DNAwas stored at -20 °C.hse-R:d(pGGGGACCTATCCATTCTAGAACATTCTAGAAGCTTCGAGAAGGTAGCCTCA)hse-Y:d(pCCCCTGAGCCTACCTTCTCGAAGCTTCTAGAATGTTCTAGAATGGATAGGT)hse-R•hse-Y:*** *** *** *** *** ***GGGGACCTATCCATTCTAGAACATTCTAGAAGCTTCGAGAAGGTAGGCTCATGGATAGGTAAGATCTTGTAAGATCTTCGAAGCTCTTCCATCCGAGTCCCChse-A:d(pCAAGACCTATCCATTCTAGAACATTCTAGAAGCTTCGAGAAGGTAGGCTCA)hse-T:d(pCTTGTGAGCCTACCTTCTCGAAGCTTCTAGAATGTTCTAGAATGGATAGGT)hse-A•hse-T:*** *** *** *** *** ***CAACACCTATCCATTCTAGAACATTCTACAACCTTCCACAAGGTACCCTCATGGATAGGTAAGATC TTGTAAGATCTTCGAAGCTC TTCCATCCGAGTGTTCFigure 3. DNA sequences of the oligonucleotides used to form HSE probes.Asterisks indicate the location of six consecutive NC A A N pentamers (or the complement), a variable numberof which constitute the revised HSE consensus (Amin et a!., 1988; Xiao & Lis, 1988). Phosphorylated andnonphosphorylated versions of the hse-R and hse-Y oligonucleotides were synthesized and PAGE-purified byNew England Biolabs Ltd. The phosphorylated hse-A and hse-T oligonucleotides were synthesized, and nonnucleotidic contaminants were removed from the unpurified crude products, by Oligos Etc. Inc.Assuming an average Mr of 650 per nucleotide pair, 1 .tg of the 51-bp HSE duplex is approximately30 picomoles. I also assumed that after the annealing reaction, the DNA was quantitatively precipitated(i.e., because the DNA concentration was very high); therefore I estimated that one A260 unit of the 5 1-bp HSEduplex was about 35 j.tg, based on the UV-ABS data obtained with the 2.0 mg/mL solutions.552. Fill-in end-labeling of the HSE probe.The following reagents were added (in order) to a microcentrifuge tube: 10.0 JIL ofa-[32P]-dCTP(3 jiM, 3000 Ci/mmol, 10 Ci/L), 2.0 pL of 5x Sequenase buffer, 1.0 iL of 0.1 M DTT, 1.0 pL of 2 mglmL BSA,and 1.0 iL of the duplex HSE probe (600 ng/j.iL). The reaction was pre-incubated 5 mm on ice, and then 1.0 iLof 13U/j.tL Sequenase version 2.0 T7 DNA polymerase was added. The reaction was mixed gently, thenincubated 5 mm on ice, 5 mm at 20-22 °C, and 15 mm at 37 °C (Sambrook et al., 1989). Two microlitres ofchase mix (10 mM each of dATP, dCTP, dGTP, and dTTP in 50 mM NaCl) was added, and the reaction wasincubated for a further 15 mm at 37 °C. The reaction was stopped with 2.0 iL of 0.10 MNa2EDTA pH 7.6, andthen 380 j.iL of NENSORB REAGENT A was added.A NENSORB-20 nucleic acid purification column was carefully packed with the dry resin and thenwashed with 3 mL of methanol. Any resin that did not pack into the column proper (i.e., which adhered to thewalls of buffer reservoir after the methanol was added) was dislodged and allowed to settle into the column byscraping the inner surface of the reservoir with a 1-mL pipette tip. After most of the methanol (-2 mL) haddrained through the column, the column bed was stirred gently with an ultramicropipettor pipette tip to removeair bubbles. The column was then washed with 3 mL of NENSORB REAGENT A. The labeling reaction wasloaded on and allowed to run into the column, and then the column was washed with 3 mL of NENSORBREAGENT A and 3 mL of TE pH 7.4. The probe was eluted with 1 mL of a 1:1 v:v mixture of EtOH:TEpH 7.4. Four microcenthfuge tubes were used to collect fractions (drops 1-3, 4-16, 17-24, and 25+). Thesecond fraction (drops 4-16) contained >80% of the radioactivity that had previously been retained by thecolumn. This fraction (about 250 j.tL) was precipitated overnight at 4 °C with 0.06 vol (15 i.IL) of 3.0 M NaOAc,pH 5.2 and 2 vol (600 jiL) of EtOll. After the DNA was microcentrifuged (see Topic B of this chapter) therewas no visible pellet whatsoever (the NENSORB-20 column is one of the few minicolumns that do not shed anyresin fines), and very little of the radioactivity was detectable in the supernate. The dried probe DNA wassubsequently resuspended with 400 jiL of TE pH 7.4, stored at 4 °C, and used within 10 days. The probeconcentration was estimated to be 1.0 ng/jiL, assuming an overall recovery of 67%.56Some of the EMSA results reported in this thesis were obtained with probes that were labeled usingearlier, less efficient versions of the above protocol.3. Preparation and PAGE analysis of EMSA binding reactions.The following protocol describes the approach that I used to detect HSE-specific DNA-binding activity.This approach employs electrophoresis buffers and associated running conditions that differ markedly fromeither the high-glycine-concentration electrophoresis buffer described by Schneider et al. (1986) or theconventional, low ionic-strength electrophoresis buffers that are more commonly used for EMSAs (e.g., Fried &Crothers, 1981; Strauss & Varshavsky, 1984).A 4x stock solution of acrylamides was prepared, stored at 4 °C shielded from light, and used withinone week. Polymerization of a degassed, 40-mL PAGE gel solution (3.5 to 4.5% T; 2% CB1; either KGBM,KGB H, or KGTH buffer) was initiated with 100 jiL of 30% w/v ammonium persulfate and 30 j.LL of TEMED.The solution was mixed gently but thoroughly, then poured into a 1.5-mm-thick, 18 x 16 cm gel-moldassemblyt. The polymerized gel was stored in a Zip-Loc plastic bag and left to set overnight at 20-22 °C.An aliquot of nuclear extract was thawed at 37 °C for 40 seconds, microcentrifuged for 1 mm at20-22 °C (to remove air bubbles), mixed gently, and then used immediately. The binding reactions were carriedout at 20-22 °C in 0.5-mL microcentrifuge tubes. Reaction conditions are summarized in the figure legends.After the 30-mm binding reaction was over, 9 iL of each reaction was analyzed by PAGE, which separated thefree probe (Rf=1.0) from the putative ceHSBF-HSE complex (Rf as indicated). There was no loading (ortracking) dye present in the reactions.The electrophoresis apparatus (Hoefer Scientific Instruments, model SE 600) was set up using 800 and650 mL of buffer (either KGBM, KGBH, or KGTH) in the lower and upper buffer reservoirs, respectively.Electrophoresis was carried out at 20-22 °C. The same buffer that was used in the gel was used in the runningbuffer. The electrophoresis buffer was not recirculated (lx KGBM, KGB H, and KGTH are all highly buffered)and cooling of the electrophoresis unit (i.e., running cold water through the unit’s cooling system) was nott Note that in some cases EMSA gels (whose dimensions differed from those given above) were run in anelecirophoresis apparatus which is not commercially available and is not described here.57necessary. The electrophoresis buffer should not be re-used. Typically the gel was prerun for 30 mm at 40 V(=2.5 V/cm), loaded, and then run for 6 hours at 40 V. Under these conditions the use of either KGBM, KGBH,or KGTH does not result in any heating of the gel. The gel was transferred to a piece of heavyweight art paper(Bristol board or the equivalent), and then the (unwrapped) gel was dried in a 65-70 °C oven for 2-3 hours. Thedried PAGE gel was covered with a very thin sheet of parchment paper and then autoradiographed as indicatedin the figure legends.I did not “fix” the gel prior to drying. I found that the use of a vacuum gel dryer resulted in asignificant loss of radioactivity through the (unfixed) EMSA gel (and the backing paper), presumably because ofthe small size of the DNA probe, and possibly also because of the porosity of the gel.58CHAPTER 1.3. RESULTSA. PREPARATION OF NUCLEI FROM C. ELEGANS EGGS.The A-protocol for isolating nuclei from C. elegans eggs was refined over time to maximize the yieldof nuclei while maintaining conditions designed to preserve native chromatin structuret . An earlier version ofthis procedure is the X-protocol. Nuclear extracts prepared using these two protocols yielded significantlydifferent EMSA results (see below). However, both protocols shared several important features. In both cases acell-free homogenate was prepared using a buffer that contained KC1 and MgC12 (75 and 2.5 mM in theX-protocol, and 120 and 3.5 mM in the A-protocol, respectively), 1.0 mM EGTA, and 0.4% w/v n-octyl-J3-D-glucoside (OG). In both protocols the homogenate was diluted with a buffer that was very similar to thehomogenization buffer (but with only 0.02% w/v OG), debris was cleared by low-speed centrifugation, and thennuclei present in the supernatant fraction were pelleted, washed, and repelleted. In the X-protocol the nucleiwere washed a second time prior to the extraction step, but this only appeared to reduce the yield of nuclei. Thissecond wash step was subsequently judged to be unnecessary and was omitted from the A-protocol. In bothprotocols DNA-binding proteins were extracted from the purified nuclei using 350 mM KC1 (in a final volumeof about 0.5-0.6 of the initial volume of tissue), and EDTA was used to chelate divalent cations during theextraction. In both protocols the use of a stainless steel Dounce homogenizer was required to generate enoughforce to disrupt C. elegans eggs. By comparison, glass or glass/Teflon homogenizers were very ineffective,presumably because only the stainless steel homogenizer had a clearance that was small enough to disrupt theeggs.Other parameters which did not appear to qualitatively affect the nuclei or the EMSA results were theexact concentrations of DTT (between 1 and 10 mM) and OG (between 0.22 and 0.42% w/v) in thehomogenization buffer (X.40 or A.42). However, the values given for these and other parameters in theA-protocol gave better yields of nuclei and higher levels of ceHSBFm activity.t Although chromatin structure was never examined in this study, the above statement is based primarily on thecomposition of the nuclei isolation buffers that were used (especially for the A-protocol) and the importanceascribed to these parameters by the work of Walker & Silcorska (1986).59Vanfleteren et al. (1979) reported that storage of nematodes at -80 °C softened the cuticle. I found thatthis was also true for egg tissue stored at -70 °C. The degree of softening varies from sample to sample, but ingeneral, the longer the eggs were stored at -70 °C, the easier it was to homogenize them. Therefore whateverhomogenization procedure is adopted, it is important to use microscopy to monitor disruption of the tissue, andmake appropriate adjustments regarding the degree of homogenization required.The yield of A-protocol nuclei was determined using the DNA-specific fluorochrome Hoescht 33258(Labarca & Paigen, 1980). Measured aliquots of eggs and nuclei were digested with proteinase K in thepresence of EDTA and SDS. Measured aliquots of these digests were then assayed using calf thymus DNA(42% G+C) as a standard. The apparent relative yield of A-protocol nuclei was 33 ± 1%, and the apparentabsolute yield was 0.91 ± 0.03 mg of DNA per gram of wet tissue (n=2). Using the above procedure the DNAcontent of 1 gram of eggs was estimated to be 2.8 ± 0.1 mg (n=2).The reported yields (from asynchronous cultures of C. elegans) of milligrams of DNA per gram of wettissue range from 0.6 (Vanfleteren, 1982) to 1-2 (Sulston & Brenner, 1974). Dixon et al. (1990) reported ayield of 20-40 A260 units (in 0.1 M NaOH) of crude nuclei per gram of frozen worm or egg droplets. If all ofthis A260 absorbing material was DNA, and assuming that the droplets were 67% w/v tissue (Dixon et al.,1990), this would represent a yield of 1.5 to 3.0 mg of DNA per gram of wet tissue (assuming 50 jig of DNA perA260 unit; Sambrook et al., 1989).The low yield of the procedure reported herein reflects the general problem of using conditions that areharsh enough to disrupt the chitinous shell of the eggs, but mild enough to preserve the condition of the nuclei.The purity and morphology of A-protocol nuclei were examined by differential interference contrast (DIC) andepi-fluorescence microscopy. The bulk of the nuclei were present in small clumps, and there was little visibleevidence of contamination with debris (Fig. 4). Individual nuclei showed satisfactory morphology whenexamined at high magnification (Fig. 5), and their overall purity and morphology compare favourably withsimilar data presented in previous reports (Dixon et al., 1990; Meheus & Vanfleteren, 1985).60AFigure 4. Overall purity of C. elegans nuclei isolated using the A-protocol.DAPI-stained nuclei were examined by DIC (upper panel) or epi-fluorescence (lower panel) methods using aZeiss IM-35 microscope. Both the field of observation and its orientation are the same in both panels.61I0.01mmFigure 5. Morphology of C. elegans nuclei isolated using the A-protocol.DAPI-stained nuclei were examined by DIC (upper panel) or epi-fluorescence (lower panel) methods using aZeiss IM-35 microscope. Both the field of observation and its orientation are the same in both panels.62B. ELECTROPHORETIC MOBILITY-SHIFT ASSAYS (EMSAs).1. The X-protocol results: Detection of a putative ceHSBF0activity.My initial attempts to use nuclear extracts—as well as a concentrated, 350 mM KC1 “whole-egg”extract (not described)—to detect HSE-specific DNA-binding activity were unsuccessful. These attemptsinvolved the use of many different protocols for preparing the extracts, as well as testing every commonly usedEMSA electrophoresis buffer system. These difficulties led me to modify (i) the conditions used to prepare thenuclei, and (ii) the electrophoresis buffer and associated running conditions used for the EMSAs.I later realized that my initial attempts to detect HSE-specific DNA-binding activity were doomed fromthe outset. During the course of this work the original definition of the HSE consensus sequence was shown tobe incorrect (Amin et al., 1988; Perisic et al., 1989; Xiao & Lis, 1988). The original “HSE” probes that I usedcontained the obsolete “Pelham” (1982) consensus sequence I G G A A T N T T C T A A as indicatedin Fig. 6. These probes lacked the trimeric sequence N G A A N N T T C N N GA A N which is nowbelieved to constitute the HSE consensus (but see Fernandes et al., 1994).** *** *** ** ** *** *** **CGCTAGCGAGCAGCTCGAATGTTCTAGCGAGCAGCTCGAATGTTCTAGAGTCGACCGGGCCCCGGGCGATCGCTCGTCGAGCTTACAAGATCGCTCGTCGAGCTTACAAGATCTCAGCTGGCN----N X----XNhe I Unequal crossover site Xba IFigure 6. The “h2” Apa I fragment bearing two copies of the obsolete HSE consensus sequence.Asterisks indicate the location of two copies of the obsolete HSE consensus. This 51 -bp Apa I fragmentcontained sequences from the intergenic region in the C. elegans hspl6-1, hspl6-48 gene pair (see Candido etal., 1989). I constructed a specialized vector for unequal, recombinant crossing over (e.g., see Rosenfeld &Kelly, 1986), and molecular clones carrying 2, 4, 8, 16, or 32 copies of the obsolete HSE consensus sequencewere obtained using the isocaudemeric Nhe I and Xba I sites as indicated. Unfortunately, the third N G A A Npentamer (i.e., in the revised HSE consensus) was replaced by vector sequences at the Xba I cloning site.Restriction sites outside of the Apa I sites were used to obtain “HSE” probes (and for fill-in end-labeling).When a duplex oligonucleotide probe bearing two thmers of the revised HSE consensus (see Fig. 3)was used in EMSAs with the “whole-egg” extract, no HSE-specific DNA-binding was observed (data notshown). When this same probe was used with an X-protocol, heat-shock nuclear extract, the EMSA results wereambiguous (see Fig. 7). Two types of putative ceHSBF activity were detected initially (Fig. 7a). One type oflal Uncertain specificityBSA Mg KHGIuHNHNHNI b IHSE-speciflcRf Extract +FWNH63Figure 7. EMSA detection of a putative ceHSBF0activity.Aliquots of the same X-protocol heat-shock nuclear extract (1.0 mglrnL of 0.35 M KC1-extracted nuclearprotein) were used in standard binding reactions {30 mm at 20-22 °C, 10 I.tl final vol, KGBH as the bindingreaction buffer, 6% wlv sucrose, 1 ng=30 fmol of a[32P]-labeled HSE probe per reaction, in both (a) and (b)}.Additional KC1 was present as indicated by the amount of extract added. KGBH is 100 mM KHG1u, 20 mMBIS-TRIS, 20mM l{EPES, pH 7.0. PAGE (using KGBH) separated the free probe (F; Rf=1.0, indicated on theleft) from the less-mobile complex (Rf as indicated). Gels were 4.5%T, 2.2%CBj8 (a) or 4.0%T, 2.O%CBj8 (b),and were prerun 1.5 h and run 4 h at 100 V (a) or 50 mA (b). Dried gels were autoradiographed for 40 h (a) or20 h (b) at -70 °C using two intensifying screens. Nonspecific competitor (nsc) DNA (N) was Hae ifi-cut DNAfrom M. luteus. Mass-excess of competitor DNA is relative to the[32Pj-labeled HSE probe.(a) Uncertain specificity. All reactions contained 1 j.tL of (b) HSE-specific binding. All reactionsextract and a 500-fold mass-excess of nsc DNA. contained 10 .tg each of tRNA and pcJN5.Reactions 2-4 were supplemented with eitherwater (W), a 550-fold mass-excess of nsc DNA(N), or a combination of a 50-fold mass-excess ofunlabeled HSE probe DNA and a 500-fold mass-excess of nsc DNA (H).At e=0 2 .tL of extract was added to reactions 2-4.Rf WHN12 3456781234Reactions were supplemented with either water (W), BSAto 0.10 mg/mL (BSA), MgCl to 5 mM (Mg), or KGBH togive total, final concentrations of 200 mM KHG1u, 40 mMBIS-TRIS, 40 mM HEPES, pH 7.0 (KHGIu).At t=0 reactions were supplemented with a 50-fold mass-excess of either unlabeled probe (H) or nsc DNA (N).64activity was designated ceHSBFm, as the protein-DNA complex (Rf of .-0.4 to 0.5) fonned by it and the probemigrated neai the middle of the EMSA gel. For the purposes of discussion, this signal will be referred to as the“Rf 0.4” signal, but in some EMSAs this signal appeared to be resolvable into a closely spaced doublet (see Figs.11 and 23a). The Rf 0.4 signal (Fig. 7a, even-numbered lanes) attributed to the [ceHSBFm—HSE probe]complex was very weak. However, the ceHSBF activity did appear to be sequence-specific, for whenever a50-fold mass-excess of unlabeled HSE probe was present (Fig. 7a, odd-numbered lanes) the Rf 0.4 signal wasdecreased substantially.The second type of putative ceHSBF activity that was detected in the X-protocol heat-shock nuclearextract was designated ceHSBF0,as the protein-DNA complex (Rf of —0.1 to 0.2) formed by it and the probemigrated near the origin of the EMSA gel (Fig. 7a, b). For the purposes of discussion, this signal will bereferred to as the “Rf 0.1” signal, but it should be noted that in some cases (e.g., Figs. 7a, 7b, and 11) this signalappeared to be attributable to the formation of at least two distinct complexes, with Rf values of 0.09 and 0.14.The Rf 0.1 signal (Fig. 7a) attributed to the [ceHSBF0— SE probe] complex(es) was much stronger that the Rf0.4 signal. However, it was not clear if the putative ceHSBF0activity was sequence-specific. If a 50-fold mass-excess of unlabeled HSE probe was present in the binding reaction, the Rf 0.1 signal was either unaffected (Fig.7a, lanes 1 and 3) or decreased only moderately (Fig. 7a, lanes 5 and 7). Although the presence of 0.10 mg/mLBSA in the binding reaction had no effect, the presence of 5 mM MgCl2 or a final concentration of 200 mMKHG1u in the binding reaction did increase the ability of the unlabeled HSE probe DNA to compete with thelabeled probe for binding with the putative ceHSBF0activity (Fig. 7a, lanes 3-8).The EMSA results shown in Fig. 7b illustrate the difficulty in determining whether or not the so-called‘ceHSBF0”DNA-binding activity was in fact sequence-specific. All of the binding reactions in these EMS Ascontained 10 p.g each of tRNA and pdN5 (random-sequence pentanucleotides) (see Zimarino & Wu, 1987). Thepresence of a 10,000-fold mass-excess of one or both of these components in the binding reaction abolished theRf 0.4 signal but had relatively little effect on the Rf 0.1 signal (Fig. 7b). The presence of a 550-fold mass-excess of nonspecific competitor DNA in the binding reaction had no effect on the Rf 0.1 signal (Fig. 7b, lane 3).65Figure 8. The influence of competitornucleic acids and heparin on the putativeceHSBF0activity.Standard binding reactions were used (seeFig. 7), except that 10 .tg of tRNA, 10 .ig ofpdN5, and 0.5 ng=15 fmol of labeled HSEprobe were present in each reaction.Reaction 1 contained the free probe (F).The same nuclear extract used in Fig 7 wasadded to reactions 2-10 (2 .tL each). Att=0, reactions 2-10 were supplemented witheither water (W) or the indicated mass-excess of either: a purified 5 1-bp restrictionfragment with two copies of the obsoleteHSE consensus sequence (h2) (see Fig. 6); apurified 681-bp restriction fragment with 32copies of the obsolete HSE consensussequence (h32); unlabeled HSE probe DNA(H); poly(dI-dC)•poly(dI-dC) (Lot I-i)(Ni); heparin (N2); C. perfringens DNA(N3); M. luteus DNA (N4); and Hae Ill-cutDNA from M. luteus (N5). PAGE andautoradiography (40 h) were performed asdescribed for Fig. 7b except the gel was3.5%T, 2%CB1S.However, addition of a 50-fold mass-excess of unlabeled HSE probe and a 500-fold mass-excess of nonspecificcompetitor DNA to the binding reaction abolished the Rf 0.1 signal (Fig. 7b, lanes 2-4).A subtle difference between the binding reaction conditions employed in the Fig. 7a and 7b EMSAswas the order of addition of reaction components. In the Fig. 7b EMSAs’ binding reactions the nuclear extractwas added last, whereas in the Fig. 7a reactions the competitor DNA was added last. I realized later that theorder of addition of the unlabeled HSE probe to the binding reaction was particularly important: if it is addedbefore the labeled HSE probe is mixed with the nuclear extract, then any observable competitive effect (i.e.,signal reduction) tends to be greater than if the unlabeled HSE probe is added last. Depending on theexperiment, the last component added to the binding reaction should therefore be the nuclear extract or thelabeled HSE probe.R 20-fold 500-foldF W h2 h32 H Ni N2 N3 N4 N5oi 2 3 4 5 6 7 8 9 1066Mother series of EMSAs compared competitor nucleic acids bearing copies of the obsolete (see Fig. 6)or the revised HSE consensus sequence for their effect on the putative ceHSBF0activity (Fig. 8). The presenceof a 20-fold mass-excess of either the “h2” or “h32” fragments in the binding reaction had relatively little effect,whereas a 20-fold mass-excess of the unlabeled HSE probe substantially decreased the Rf 0.1 signal (Fig. 8,lanes 3-5).The DNA of C. elegans is 36% (3-i-C (Suiston & Brenner, 1974). Nonspecific competitor nucleic acidsof differing base composition were therefore examined for their effect on the putative ceHSBF0activity (Fig. 8).The presence of a 500-fold mass-excess of DNA of low (26%; from C. perfringens) or high (72%; fromM. luteus) G+C content (Fig. 8, lanes 8 and 9) in the binding reaction had relatively little effect on the Rf 0.1signal. In addition, the size of the nonspecific competitor DNA did not affect its ability to compete with thelabeled probe for binding with the putative ceHSBF0activity (Fig. 8, lanes 9 and 10). As reported previouslyfor HSF and in other systems, the synthetic, alternating copolymer poly(dI-dC).poly(dI-dC) (Fig. 8, lane 6)behaved similarly to natural DNAs as a nonspecific competitor (Carthew et at., 1985; Kingston et at., 1987;Larson et at., 1988; Mosser et at., 1990; Singh et at., 1986; Sorger et at., 1987).The presence of a 500-fold mass-excess of the nonspecific competitor heparin (a polysulfatedglycosaminoglycan) substantially decreased the Rf 0.1 signal (Fig. 8, lane 7). Various HSBF isolates have beenshown to bind to heparin under the conditions (—170mM total K) used in this EMSA binding reaction (CarmoAvides et a!., 1990; Czarnecka et at., 1990; Goldenberg et at., 1988; Sorger & Petham, 1987; Wiederrecht et al.,1987; Wu, 1985; Wu et at., 1987).The presence of nonionic detergents in the binding reaction had very little effect on the Rf 0.1 signal(Fig. 9). Triton X-100 (a mixture of nona- and deca-[ethyleneglycol]-p-tert-octylphenyl ethers) had no negativeeffect (and may have even enhanced the Rf 0.1 signal slightly), even at a final concentration of 1% w/v in thebinding reaction (Fig. 9, lanes 8-1 1). n-Octyl-3-D-glucopyranoside also had little, if any, negative effect up to afinal concentration of 0.5% w/v in the binding reaction, but did appear to have a negative effect at 1.0% w/v(Fig. 9, lanes 4-7). Most HSF isolates have been shown to tolerate moderate concentrations (to —2% w/v) ofvarious nonionic detergents in EMSA and chromatographic binding reactions (e.g., see Mosser et at., 1990).67R1 OG Tx-100F W H 0.04 0.1 0.5 1.0 0.04 0.1 0.5 1.0Figure 9. Tolerance of nonionic 0detergents by the putativeceHSBF0activity.Standard binding reactions were 0.1used (see Fig. 7), except that 10ig of tRNA, 10 jig of pdN5, 250ng of Hae 111-cut DNA from M.luteus, and 0.5 ng=15 fmol oflabeled HSE probe were present ineach reaction. Reaction 1contained the free probe (F). Thesame nuclear extract used in Fig 7was added to reactions 2-11 (2 jiLeach). At t=0, reactions 2-11 weresupplemented with either water(W), a 20-fold mass-excess ofunlabeled HSE probe DNA (H),or either n-octyl glucoside (OG)or Tnton X-100 (Tx-lOU) to givethe indicated final concentration(% wlv). PAGE and autoradiography (40 h) were as describedfor Fig. 7b except the gel was3.5%T, 2%CBIS.2. The AP-protocol results: Detection of both types of putative ceHSBF activity.The main difference between the AP-protocol and the X-protocol was the composition of the buffersused during homogenization of the eggs and washing of the crude nuclei. The AP-protocol (and the A-protocol)buffers were formulated in order to preserve native chromatin structure during the purification of the nuclei (e.g.,see Murdoch eta!., 1991; Walker & Silcorska, 1986). These buffers contained 120mM KC1 and 3.5 mMMgC12,whereas the same salts were present at only 75 and 2.5 mM, respectively, in the corresponding buffers in theX-protocol. Mother buffer ingredient whose concentration may have been important was PMSF. This proteaseinhibitor was present at 0.4 and 0.2 mM in the X-protocol buffers X.42 and Y.02, respectively, but at only0.1 mM in the corresponding buffers in the AP-protocol.1 2 3 4 5 6 7 8 9 10 1168The EMSA results obtained using an AP-protocol nuclear extract prepared from heat-shocked eggswere quite different from the results obtained with the X-protocol nuclear extract. The AP-protocol extractshowed much greater levels of the putative ceHSBFm activity, whereas this activity was very difficult to detectin the X-protocol extract (cf. Fig. 10 and Fig. 7a). As shown in Fig. 10, the putative ceHSBFm activity had anapparent sensitivity to the presence of a 10,000-fold mass-excess of tRNAt in the binding reaction, but wasrelatively unaffected by the same mass-excess of pdN5 or a 1,000-fold mass-excess of poly(dI-dC).poly(dI-dC).Figure 10. Enhanced detectionof a putative ceHSBFm activity.An AP-protocol nuclear extract(prepared using heat-shockedeggs; 0.9 mg/mL of 0.35 MKC1-extracted nuclear protein)was used in standard bindingreactions (see Fig. 7). Reaction1 contained the free probe (F).Nuclear extract was added toreactions 2-10 (2 iL each). Att=0, reactions 2-10 weresupplemented with either water(W), or the indicated mass-excess of one or more of thefollowing: tRNA (T: 10,000-fold); pdN5 (P: 10,000-fold);poly(dI-dC)’poly(dI-dC) (N:Lot I-1;1,000-fold); unlabeledHSE probe DNA (USE: 50-fold). PAGE and autoradiography (15 h) were as describedin Fig. 7b except the gel wasprerun0.Shandrun6h.t It should be noted that the putative ceHSBFm activity had an apparent sensitivity to a 10,00-fold mass-excessof a particular preparation of yeast tRNA. No investigation was made to determine if tRNA itself or some othercomponent of this tRNA preparation was actually responsible for the observed inhibition of protein-DNAcomplex formation, nor was the effect of using lower concentrations of tRNA in the binding reaction examined.pF WT T T T Tp p p pN N N N NHSEFigure 11. The influence of KC1concentration on the two types ofputative ceHSBF activity.Standard binding reactions wereused (see Fig. 7), except that thebinding reaction buffer was 20 mMBIS-TRIS, 20 mM JIEPES, pH 7.0,with all of the KC1 supplied by theextract and supplementary KC1. Thesame nuclear extract used in Fig 10(1.5 j.tL) and 100 ng of pdN5 werepresent in each reaction. At t=0,supplementary KC1 was added toreactions 2-10 to give the indicatedfinal concentration (mM), which inall cases included —50 mM KC1supplied by the extract. PAGE wasas described in Fig. Th except the gelwas prerun 0.5 h and run 5 h. Thedried PAGE gel was autoradiographed for 2 h at -70 °C using oneintensifying screen.69The behaviour of the putative ceHSBF0activity was unusual, and was possibly indicative of instabilityof this activity in the AP-protocol nuclear extract. The Rf 0.1 signal was even more diffuse than usual, exceptwhen a 10,000-fold mass-excess of tRNA (or both tRNA and pdN5) was present in the binding reaction (Fig. 10,lanes 3 and 6). The Rf 0.1 signal was also uncharacteristically and erratically sensitive to the presence of a1,000-fold mass-excess of poly(dI-dC)•poly(dI-dC) in the binding reaction (Fig. 10, lanes 5 and 7-9).The AP-protocol, heat-shock nuclear extract was used in a series of EMSAs to determine the KC1concentration optima for binding of the two types of putative ceHSBF activity to the HSE probe (Fig. 11).Transfer RNA was not added to these EMSA binding reactions because of its effect on the putative ceHSBFmactivity. The KC1 concentration optimum for the putative ceHSBFm activity (—150-175 mM) was within thephysiological range. However, the value of the same parameter for the putative ceHSBF0activity (—250-350mM) was unexpectedly high.R [KCI] in the binding reaction (mM)50 75 100 125 150 175 200 250 300 3500 w’0.41.01 2 3 4 5 6 7 8 9 103. The A-protocol results: Detection of a putative ceHSBFm activity.The A-protocol is a simplified version of the AP-protocol. The unsupplemented buffers in eachprotocol were the same, but some of the buffer supplements differed. PMSF was present (at 0.10 mM) in theaElectrophoresisatpH6.3I bIElectrophoresisatpH7.9HSextractCNTextractHSextractCNTextractR[KCI]in100150thebindingreaction(mM)200250100150200250Rf[KCI]in100150thebindingreaction(mM)2002501001502002500—i’-—0--.0.41.0Ww0.5__123456781234567871Figure 12. The influence of electrophoresis buffer pH on the detection of the putative ceHSBFm activity.A-protocol nuclear extracts, prepared using either heat-shocked or control eggs (HS or CNT; 0.9 or 1.0 mglmLof 0.35 M KC1-extracted nuclear protein, respectively), were used in standard binding reactions (see Fig. 7)except that 2 tg of pdN6—but no KGBH binding reaction buffer—was present in each reaction. The extract(2.8 .tL per reaction) supplied sufficient pH-buffering and —100 mM KC1. The final concentration (mM) ofKCI present in each reaction is indicated. Supplementary KCI was added last. PAGE gels were 4.5%T,2.O%CBis and were prerun 0.5 h and run 6 h at 40 V. The dried PAGE gels were autoradiographed together forsix hours on the same piece of X-ray film (without screens).The following solutions were used for the PAGE gel and electrophoresis running buffer:(a) KGBM: 150mM KHG1u, 50mM BIS-TRIS, 50mM MES (pH 6.3);(b) KGTH: 150mM KHG1u, 50mM TRIS, 50 mM HEPES (pH 7.9).AP-protocol buffers (except buffer B), but it was not used in the A-protocol. Conversely, leupeptin and bestatmwere present (at 10 jiM each) in the A-protocol buffers (except buffer B), but were not used in the AP-protocol.There were also differences in the concentrations of OG and DT in the buffers used in each protocol (see Table2), but these differences did not appear to affect the relative abundance of the two types of putative ceHSBFactivity.Figure 13. The apparent sequence- R I ‘ ‘- ‘‘specificity oftheputativeceHSBFrn f B HSE P0 poly(dI-dC)activity. 0 50 50 100 200 500 1000 2000An A-protocol nuclear extract (prepared ousing heat-shocked eggs) was used instandard binding reactions (see Fig. 7)except that 2 jig ofpdN6—but no KGBHbinding reaction buffer—was present ineach reaction. The extract (4.0 jiL perreaction; added last) supplied sufficientpH-buffering and —140 mM KC1. Allreactions included 1 jiL of either KBHEbuffer (B), or the indicated mass-excessof unlabeled HSE probe DNA (HSE) orpoly(dI-dC).poly(dI-dC) (Lot 1-2)prepared as a lOx stock solution inKBHE. KBHE is 100 mM KCI, 20 mMBIS-TRIS, 20 mM HEPES, 1 mM EDTA,pH 7.0. The final, total concentration ofKC1 was —150 mM in each reaction.PAGE was performed using KGBM as 1.0described in Fig. 12. The dried PAGE gelwas autoradiographed for two hours at-70 °C using one intensifying screen.0.4 41 2 3 4 5 6 7 872The putative ceHSBF0 activity could not be detected in A-protocol nuclear extracts prepared fromheat-shocked or control eggs (Fig. 12). The putative ceHSBFm activity was readily detected in A-protocol,heat-shock nuclear extracts, but appeared to be slightly less abundant in control extracts prepared using the sameprotocol. In most experiments the Rf 0.4 signal was stronger and sharper when PAGE was performed using aslightly acidic (—pH 6.3) electrophoresis buffer (Fig. 12a). This particular buffer system (KGBM) worked verywell, but it should be stressed that the Rf 0.4-0.5 signal attributed to the putative ceHSBFm activity could bedetected using KHG1u-based electrophoresis buffers at pH 7.0 and 7.9 also (Figs. 7a, 10, 11, and 12b).Figure 14. Order-of-addition analysisof ceHSBFm sequence-specificity.Parts (b) and (c) of this figure arelocated on the next page. Standardbinding reactions were used (see Fig.________________________________12) except as noted. The same heat-shock extract used in Fig. 12 (4.0 pL)was present in each reaction, and itsupplied sufficient pH-buffering and-440 mM KC1. In each set of EMSAs(a,b, or c) the indicated amount of[32P]-labeled HSE probe was presentin each reaction. AU reactions included1 j.tL of either KBHE buffer (B), or theindicated mass-excess of competitorDNA (comp.)—unlabeled HSE probe(HSE) or poly(dI-dC).poly(dI-dC) (Lot1-2) (nsc)—prepared as a lOx stocksolution in KBHE. Each reaction hadthe indicated component added last.The final, total concentration of KC1was —150 mM in each reaction. PAGEwas performed using KGBM asdescribed in Fig. 12. The dried PAGEgels were autoradiographed for 6 h (a;without screens) or for 7 h (b) or 80 h(c) at -70 °C using one intensifyingscreen.I aj 30 fmol of probe par reactionAdded last to the binding reaction:HS extract probe comp.Rf B nsc HSE nsc HSE0 2000 50 2000 500 — — —nsc HSE2000 501 2 3 4 5 6 7j bI3fmolofprobeperreactionICj300amolofprobeperreactionAddedlasttothebindingreaction:I-ISextractprobecomp.Addedlasttothebindingreaction:HSextractprobecomp.RBnscHSEnscHSEnscHSE02x104502x104502x10450,.%it.”—R1BnscHSEnscHSEnscHSE02x10502x105502x105500‘‘i—0.40.4 1.0-1.01234567123456774The putative ceHSBFm activity present in A-protocol heat-shock nuclear extracts did appear to be veryspecific for the HSE probe (Fig. 13). A-protocol control nuclear extracts gave identical results (data not shown).An order-of-addition effect was noted, a behaviour similar to that displayed in previous EMSAs by the putativeceHSBF0activity. This effect was examined using various amounts of labeled HSE probe in the EMSA bindingreactions (Fig. 14). In these binding reactions there was no pre-incubation period (other than the time requiredfor pipetting) prior to the addition of the component that was added last to the binding reaction (whose additiondefined the start of the 30-mm incubation period for the complete reactions). In most cases the presence of a50-fold mass-excess of unlabeled HSE probe in the binding reaction substantially reduced the Rf 0.4 signal(Figs. 13-14b). However, when the amount of labeled HSE probe in each reaction was lowered to 300 amol, thepresence of a 50-fold mass-excess (15 fmol) of unlabeled HSE probe had virtually no effect on the Rf 0.4 signal,presumably because the total amount of HSE probe (labeled and unlabeled) was limiting relative to the amountof putative ceHSBFm activity in 4 iL of A-protocol heat-shock nuclear extract (Fig. 14c). In these EMSAs thepresence of a 2,000- to 200,000-fold mass-excess of poly(dI-dC)•poly(dl-dC) in the binding reaction had verylittle effect on the Rf 0.4 signal (Figs. 13-14).Figure 15. The influence of proteinase K and nonionic detergents on the putative ceHSBFm activity.This figure is located on the following page. The same extracts used in Fig. 12 were used in standard bindingreactions (as specified in Fig. 12 and described further below). PAGE was performed using KGBM asdescribed in Fig. 12. The dried PAGE gels were autoradiographed for six hours (without screens).(a) Proteinase K sensitivity. Reactions 3-6 contained 2.8 jiL of the indicated nuclear extract. Control reactionscontained 2.8 IlL of either water (W; lane 1) or a 5:1 v:v mixture (AB; lane 2) of nuclei isolation buffers A.02and B (respectively) instead of nuclear extract. Water (W; lanes 3 and 5) or proteinase K dissolved in water (P;0.010 U; lanes 4 and 6) was added last to each reaction.(b) Tolerance of nonionic detergents. All reactions contained 4 iL of the indicated nuclear extract. The fmalconcentration (%w/v) of n-octyl glucoside (OG) or Triton X-100 (Tx-100) present in each reaction is indicated.Water (W) or nonionic detergents were added last to each reaction.lalProteasesensltlvItyIbjToleranceofnon-IonicdetergentsHSextractOGTx-100W0.51.00.51.00—--imw-irExtractpresentHSwPCNTwpExtractRabsentWAB00.4RCNTextractOGTx-100W0.51.00.51.0—4_0.4___1234567891012345676The Rf 0.4 signal was not attributable to the formation of a complex between the labeled HSE probeand any of the polypeptide or nonpolypeptide protease inhibitors present in the extraction buffers (bufferA.02:buffer B, 5:1, v:v), as this signal could not be detected in the absence of salt-extracted nuclear proteins(Fig. 15a, lane 2). This signal did arise from the formation of a protein-DNA complex, as the addition ofDNase-free proteinase K to the binding reaction substantially decreased the Rf 0.4 signal (Fig. 15a, lanes 4and 6). The presence of 0.5-1.0% wlv of n-octyl glucoside or Triton X-100 in the binding reaction had either noeffect or slightly enhanced the Rf 0.4 signal, respectively (Fig. 15b). These results were very similar to thoseobserved earlier with the putative ceHSBF0activity (Fig. 9).77CHAPTER 1.4. DISCUSSIONA. THE STUDY OF SEQUENCE-SPECIFIC DNA-BINDING PROTEINS FROM C. ELEGANS.Two iranscription factor genes from C. elegans (mec-3 and unc-86) have been expressed in E. coli, anda third C. elegans gene (tra-1) has been expressed in vitro, and preliminary biochemical studies of theserecombinant proteins have been reported (Xue et al., 1992; Zarkower & Hodgkin, 1993). However, nosequence-specific DNA-binding proteins isolatedfrom C. elegans have been studied at the biochemical level,despite the interest that this species holds for developmental biologists, geneticists, molecular biologists, andneurophysiologists, and despite the caveats that must be attached to results obtained using recombinanteukaryotic transcription factors (e.g., see the above discussion of HSF).Any study of a sequence-specific DNA-binding protein isolated from C. elegans must consider thedevelopmental stage and amount of nematode tissue required for such an investigation. I chose egg tissue forthis study for several reasons. Cells at this developmental stage express heat-shock genes, and hence these cellswere expected to contain the sequence-specific DNA-binding protein under study (Snutch & Baillie, 1983).Another reason for choosing nematode eggs is that nuclei purified from this tissue source do not containsignificant protease or nuclease activities (Dixon et al., 1990; see also Fig. 23 of Part 2 of this thesis). Cell-freehomogenates prepared from larval or adult stage nematodes may contain extracellular proteases and nucleasessecreted by cells lining the digestive tract, a problem that is avoided by the use of egg tissue. It is also difficultto prepare nuclei from larval or adult stage nematodes because the cuticle is very resistant to disruption.The marn drawback to the use of nematode eggs as a starting material is the loss of valuable biomassduring their preparation. In addition, there are practical limits to the amount of material that can be processed byone person during the procedure where the gravid nematodes are exposed to an alkaline hypochlorite solution.If it is known that the sequence-specific DNA-binding protein to be isolated from C. elegans is present in eggtissue, it may be preferable to use “whole-egg” or nuclear extracts prepared from egg tissue in smaller scale,exploratory studies to determine optimal conditions for assaying the protein of interest. One may then want touse a tissue homogenate prepared from adult nematodes in larger scale preparative work. Gbewonyo et al.78(1994) have reported the large-scale culture of C. elegans, and this work is an excellent guide for the preparationof kilogram amounts of nematodes.B. THE DETECTION OF TWO TYPES OF PUTATIVE ceHSBF ACTIVITY1. Difficulties in identifying ceHSBF activity.The first and most serious difficulty that I had in identifying ceHSBF activity was selecting the DNAsequence for the HSE probes, as the initial consensus sequence proved to be incorrect (Amin et al., 1988; Pensicet al., 1989; Xiao & Lis, 1988). The discoveries by other researchers of the interspecies diversity displayed byHSF isolates were also surprising, as were the discoveries that many species possess more than one type of HSFactivity (see Sorger, 1991). The most unexpected discovery was the report of La Volpe et a!. (1988; see alsoNaclerio et a!., 1992) of the RcC9 family of repetitive elements, or “HSE islands”, in the DNA of C elegans.No other eukaryotic genome has been shown to have this bizarre structural feature. Thus even after the problemwith the HSE probes was corrected, I was still uncertain as to the number, relative abundance, electrophoreticmobility-shift behaviour, or other properties of the putative ceHSBF activity (or activities) that I might be able todetect.Some comments on the reproducibility of the EMSA results obtained with the three types of nuclearextract should also be noted. Only one preparation of each of the X-protocol and AP-protocol nuclear extractswere obtained. Thus different preparations of either of these two extracts were not compared, but differentaliquots from either the X-protocol or AP-protocol preparations yielded consistent EMSA results. However, tendifferent preparations of an A-protocol heat-shock nuclear extract, and three different preparations of anA-protocol control nuclear extract, were obtained and examined by EMSA analysis. The A-protocol nuclearextract preparations yielded consistent EMSA results, and the EMSA results obtained using different aliquotsfrom a given preparation were also consistent. Rather poor EMSA results were obtained with only onepreparation of each of the heat-shock and the control A-protocol nuclear extracts, but both of these extractswere prepared at the same time using the same batch of nuclei-isolation buffers, and the problems encounteredwere cleared up when these buffers were discarded and fresh buffers were used.79The EMSA analysis of the putative ceHSBF0 activity is therefore incomplete and particularlyunsatisfactory, as this activity was originally discounted as an artifact, and only one preparation of each of theX-protocol and AP-protocol extracts were examined. The EMSA analysis of the putative cCHSBFm activitycertainly could be improved upon, but at least it is known that the EMSA results obtained using the A-protocolextracts were very reproducible—not only for a given extract preparation, but also for each of the many extractsprepared using the A-protocol.Many different experimental approaches can provide evidence that an EMSA-detectable protein-DNAinteraction under study is target sequence-specific. In this discussion the target” refers to the cognate bindingsite of the sequence-specific DNA-binding protein of interest: let wild-type and functionally defective variantsof this sequence be denoted by “T+” and “T”, respectively. Although a given “target probe” by definitioncarries this sequence [T+], there are “flanking” sequences on either side of “T+” which may act as binding sitesfor other DNA-binding proteins: denote these flanking sequences on a particular probe as “F+”, and unrelatedflanking sequences on another probe as “F’. Given a particular target probe [a T+F+ probe], the experimentalapproaches that provide evidence that an EMSA-detectable protein-DNA interaction under study is targetsequence-specific include the use of control EMSA probes (or competitor DNAs) that are (or about) the samelength as the target probe, but with a DNA sequence that is either: (i) unrelated to the target probe [a TFprobe], or (ii) a target-specific, target-abolishing sequence variant of the target probe [a T—F+ probe], or (iii) asecond target probe with unrelated flanking DNA [a T+F_ probe]. In the case of EMSAs, four questions arerelevant to the determination of target sequence-specificity: Does the signal attributable to the formation of aprotein-DNA complex arise from binding of protein(s) to: (i) the termini of any probe; (ii) any sequence on anyprobe; (iii) a specific flanking sequence on the target probe; or (iv) the target sequence on the target probe.Probably the most rigorous EMSA evidence that a protein-DNA interaction under study is targetsequence-specific is the following: (i) the EMSA signal of interest is observed when a TF probe is used; (ii)this signal is substantially reduced when a moderate (in most cases, 50- or 100-fold) mass-excess of a T+F(unlabelled) probe is used as a specific competitor DNA; (iii) this signal is unaffected when a substantial80mass-excess (say,>> 100-fold) of a T—F+ probe is used as a “nonspecific” competitor DNA; and (iv) the signalof interest is not observed when a TF + probe is used as a probe.In the literature it is common to see the following data provided as EMSA evidence ofsequence-specificity: (i) the EMSA signal of interest is observed when a TF probe is used; (iii) this signal issubstantially reduced when a moderate (in most cases, 50- or 100-fold) mass-excess of (unlabelled) T+F+ probeis used as a specific competitor DNA; (iii) this signal is unaffected when a substantial mass-excess (say,>>100-fold) of a T—F— probe is used as a “nonspecific” competitor DNA; and (iv) the signal of interest is notobserved when T—F— probe is used as a probe. Often the EMSA evidence is supplemented with other datasupporting an assumption of target sequence-specificity, but in the above cases where a TF probe orcompetitor DNA is used as a control, the EMSA data alone does not really distinguish between target [T+j andflanking [F+J sequence-specific binding to the T+F+ probe. Indeed, there is little difference between the use ofa substantial mass-excess of a TF probe as a “nonspecific” competitor DNA and the use of random-sequenceDNA or synthetic alternating copolymer DNA such as poly(dI-dC).poly(dI-dC) for this purpose.At the beginning of this study, various ‘HSE” [“T+F+”J and unrelated [TFj control probes wereused. All of these probes were plasmid-based, and were either cloned inserts bearing one or more copies of theoriginal HSE, or unrelated DNA fragments of comparable size (e.g., 65- or 75-bp Hinf I fragments frompUC 13, or a 65-bp Nhe I—Rsa I fragment from pBR 322). When various nuclear extracts prepared fromheat-shocked nematode eggs were used in EMSAs with these probes, even the most rudimentary controls(competition with nonspecific DNA) showed that none of the EMSA signals observed could possibly beattributable to sequence-specific protein-DNA interactions. Later, when the original HSE consensus sequencewas shown to be incorrect, I viewed the so-called ‘HSE’ probes (that were based on the original consensussequence) as control probes—not totally unrelated to, but not really “target-specific, target-abolishing sequencevariants” of, the hse-R•hse-Y and hse-A•hse-T probes that I eventually did use (the plasmid-based andoligodeoxyribonucleotide probes did not have the same flanking DNA). Admittedly the nuclear extracts and theelectrophoresis buffers that were used in the early EMSAs (that used plasmid-based probes) differed from the81extracts and buffers used in the EMSAs whose results are reported herein, but in my view they were not so farremoved as to render the information derived from the early EMS As as irrelevant to subsequent work.The signals attributable to the two types of putative ceHSBF activity were both substantially reducedwhen a moderate (50-fold) mass-excess of (unlabelled) target probe was used as a specific competitor DNA, yetthese signals were unaffected when a substantial mass-excess (500-fold to 200,000-fold) of unrelated DNA wasused as a nonspecific competitor. It is therefore unlikely that the two types of putative ceHSBF activity arosefrom nonspecific protein-DNA interactions. It is also unlikely that the two types of putative ceHSBF activityarose from the binding of protein(s) to the ends of the probe. It must be remembered that when a 50-foldmass-excess of unlabelled probe was used as a specific competitor DNA, the termini of the DNA were not“filled in” and were therefore 4-nucleotide-long single-stranded regions. This was in contrast to the1-nucleotide-long single-stranded overhang on the probe, which was labeled with a modified T7 DNApolymerase that is incapable of producing flush ends (Sambrook et al., 1989). The 5’-phosphorylated andnonphosphorylated versions of the hse-R•hse-Y probes, as well as the 5’-phosphorylated hse-A•hse-T probe,(Fig. 3) could be substituted for one another without effect (both as probes and as specific competitor DNA5),and the single-stranded ends of the these probes differed. The putative ceHSBF0activity tolerated a 10,000-foldmass-excess of pdN, which is about a 100,000-fold molar-excess (relative to the labeled probe) of 5’-ends ofsingle-stranded DNA (Fig. 7b). The putative ceHSBFm activity tolerated a 200,000-fold mass-excess of pdN6,which is about a 1,600,000-fold molar-excess of 5’-ends of single-stranded DNA (Fig. 14c).Unfortunately, rigorous EMSA controls were not performed, and neither of the two types of putativeceHSBF activity were examined by DNA footprinting. Thus all that can be concluded at this point is that thesetwo DNA-binding activities appear to be specific for an HSE-bearing probe. Neither activity has beenrigorously demonstrated to be specific for the HSE consensus sequence itself.2. The putative ceHSBF0activity.In most reports of the EMSA detection of a heat-inducible HSE-DNA-binding activity, thesequence-specific protein-DNA complex displays an Rf of 0.1-0.2 when resolved by PAGE from the free probe(Rf = 1.0) (e.g., Mosser et al., 1990). The Rf value of the HSF-HSE complex is higher and more variable when82agarose is used as a gel matrix (cf. Shuey & Parker, 1986a; Zimarino & Wu, 1987). Thus even though EMSAPAGE gels are typically as porous as the matrix will allow, the electrophoretic mobility of the HSF-HSEcomplex under these conditions is primarily a function of the size and shape of the complex, not its net charge.This is understandable given the size and axial ratio of the HSF trimer (Peteranderl & Nelson, 1992; Westwood& Wu, 1993). This also explains why the reported PAGE Rf values of most HSF-HSE complexes are verysimilar, despite differences in the size of the HSE probes, the ionic strength and composition of theelectrophoresis buffers, or the percentages of acrylamide and bis-acrylamide in the EMSA PAGE gels that havebeen used in these studies.The PAGE Rf of the protein-HSE complex formed by the putative ceHSBF0activity agreed with thePAGE Rf of the corresponding complex formed by virtually all of the heat-inducible HSE-DNA-bindingactivities that have been reported in the literature. Unfortunately, a control nuclear extract from C. elegans wasnot examined for this type of putative HSBF activity, so I cannot say whether or not the HSE-binding ability ofthe putative ceHSBF0activity is heat-inducible or constitutive. A further complication to note is that though“control” eggs do not express heat-shock RNA (Jones et al., 1989), the HSE-binding ability of a ceHSF activity(i.e., a ceHSBF activity) might be induced by the stress of the procedure used to prepare the eggs (whichinvolves dissolving the gravid adults in an alkaline hypochlorite solution, a treatment which the eggs can survivefor short periods). Thus if the HSE-binding ability of a ceHSF activity can be detected in heat-shocked but notin control eggs, one can conclude that this ability is heat-inducible. However, if the HSE-binding ability of aceHSF activity can be detected in both heat-shocked and control eggs, one cannot conclude that this ability isconstitutive, for there is a possibility that it may have been inadvertently induced in the “control” eggs.It is difficult to account for the differences in the relative abundance of the putative ceHSBF0activityin the nuclear extracts prepared using the three different protocols. The relative abundance of the putativeceHSBF0 activity was inversely related to the concentration of PMSF in the homogenization and washingbuffers used in these protocols. HSF is susceptible to proteolysis fl vitro (Sorger & Petham, 1987; Wiederrechtet al., 1987; Wu et al., 1987). However, the detectable protease activity in nuclear extracts prepared fromC. elegans egg tissue is very low (Dixon et al., 1990; see also Fig. 23 of Part 2 of this thesis). In addition, the83protease inhibitors used in the A-protocol were present at adequate concentrations, and were selected on thebasis of their ability to inhibit the most prominent protease activities in this species (Sarkis et at., 1988).The EMSA analyses of nuclear extracts prepared using the three protocols reported herein were notcarried out at the same time, and the extracts themselves were prepared using eggs that had been stored at -70 °Cfor different lengths of time (with the storage period being the least and the greatest for the X-protocol and theA-protocol extracts, respectively). The putative ceHSBF0activity may not be stable to long-term storage at-70 °C, and this could account for the decline and absence of this activity in nuclear extracts prepared using theAP-protocol and its successor, the A-protocol, respectively. The putative HSF activity present in extractsprepared from the parasitic worm Schistosoma mansoni has been reported to be unstable to long term storage at-60 °C (Levy-Holtzman & Schechter, 1994). For whatever reason, the earliest of the three protocols, theX-protocol, gave the highest yield of the putative ceHSBF0activity. The recommended EMSA conditions forthe detection of this activity include: the use of 3 fmol or less of labeled HSE probe per binding reaction; thepresence of IRNA in the binding reaction at either a concentration of 1 mg/mL or at a 10,000-fold mass-excessrelative to the amount of labeled HSE probe; and the use of KGBH as the electrophoresis buffer.3. The putative ceHSBFm activity.EMSA studies using extracts prepared from mouse (Hensold et at., 1990; Mezger et at., 1989) andhuman cell lines (Abravaya et at., 1992; Lee et at., 1993; Mosser et at., 1988, 1990) have detected the formationof more than one protein-DNA complex that is specific for an HSE-bearing probe. In these reports the mostprominent probe-specific protein-DNA complex (i) is only formed when heat-shocked extracts are used, (ii)gives rise to the strongest autoradiographic signal of the complexes, and (iii) displays an Rf of -0. 1-0.2 whenresolved by PAGE from the free probe (Rf = 1.0). In the studies by Hensold et at. (1990) and Mosser et at.(1988, 1990) the Rf 0.1 complex was examined further by DNA footprinting and found to be specific for theHSE consensus sequence.The second probe-specific protein-DNA complex that was detected in these studies is formed wheneither heat-shock or control extracts are used, and displays a PAGE Rf of —0.3-0.4 (Abravaya et at., 1992;Hensold et at., 1990; Lee et at., 1993; Mezger et at., 1989; Mosser et at., 1988, 1990). This complex was not84examined further by DNA footprinting in any of these studies. Thus although it is common to refer to thesecond protein-DNA complex as “HSE-specific”, the possibility exists that some or all of them may be specificfor some other (non-HSE) sequences on the probes (e.g., flanking DNA). However, given that many species(including H. sapiens and M. muscus) have more than one HSF gene, and that considerable intraspecies diversityis displayed by these genes, the ability to detect more that one HSE-specific protein-DNA complex in thesestudies is not unreasonable.The detection in an EMSA gel of multiple bands arising from the formation of protein-DNA complexesof identical sequence-specificity may also be due to differences in protein stoichiometry, configurationalisomerism, or conformational isomerism (Fried, 1989). Fried (1989) has defined configurational isomers as‘protein-DNA complexes of identical stoichiometry, differing in the location of bound proteins” andconformational isomers as “complexes of identical stoichiometry and configuration, differing in theconformation(s) ofone or more components’The protein-DNA complexes formed by the putative ceHSBFm and ceHSBF0activities are probablyneither configurational nor conformational isomers. In most cases these types of isomers give rise to EMSAsignals whose Rf values differ only slightly (Fried, 1989). In addition, although the putative ceHSBFm andceHSBF0activities both appeared to be specific for the HSE probe, and displayed the same tolerance of OG orTriton X-100, these two types of putative ceHSBF activity showed very different behaviour when a 10,000-foldmass-excess of tRNA was present in the binding reaction (Fig. 10). It is unlikely that only one of twoconfigurational or conformational isomers would exhibit an apparent sensitivity to a 10,000-fold mass-excess oftRNA.When the amount of labeled HSE probe in the binding reaction was lowered from 30 to 3 and then to0.3 fmol, and the amount of A-protocol heat-shock nuclear extract was kept constant, there was still nodetectable ceHSBF0activity (Fig. 14). Instead there was simply an increase in the percentage of the labeledHSE probe that was complexed with the putative ceHSBFm activity. Thus the protein-DNA complex formed bythe putative ceHSBF0activity is probably not a higher order stoichiometric variant (assuming that the putativeceHSBF0activity consists of a hexamer of two HSF trimers occupying all six of the NGAAN pentamers on the85probe) of the complex formed by the putative ceHSBFm activity (assuming that the putative ceHSBFm activityconsists of a trimer of cooperatively binding 11SF monomers occupying three of the six NGAAN pentamers onthe probe). If this were so, I would have observed an increase in the amount of complex of higher orderstoichiometry (i.e., ceHSBF0)with an increase in the ratio of [nuclear extract protein] to [HSE probe DNAJ inthe EMSA binding reactions (Fried, 1989).The putative ceHSBFm activity may simply be a discrete proteolytic degradation product of theputative ceHSBF0activity. Proteolytic degradation of the putative ceHSBF0activity would be expected to giverise to one or more diffuse or smeared bands in the EMSA gel. However, in most cases the protein-DNAcomplex formed by the putative ceHSBFm activity gave rise to a single, sharp band in the EMSA gel. In somecases the Rf 0.4 band appeared to be resolved into a closely spaced doublet (e.g., Figs. 11), but this phenomenonwas not observed in other EMSAs.The observed effects of tRNA in the binding reaction also appear inconsistent with the hypothesis thatthe putative cellS BFm activity is a proteolytic derivative of the putative ceHSBF0 activity. If the putativeceHSBFm activity was a proteolytic degradation product, it is unclear how its DNA-binding ability wouldacquire its apparent sensitivity to the presence of a 10,00-fold mass-excess of tRNA, a property that its precursorlacks.If the putative ceHSBFm activity is neither a proteolytic derivative nor a stoichiometric, configurationalor conformational isomer of the putative ceHSBF0 activity, then why was there a reciprocal relationshipbetween the levels of the two activities in the X-protocol and A-protocol nuclear extracts? As discussed earlier,the lack of detectable ceHSBF0activity in the A-protocol nuclear extract may have been due to instability tostorage at -70 °C of this activity. However, the lack of detectable cellS BFm activity in the X-protocol nuclearextract was even more puzzling. One possible explanation may be that this activity was rapidly lost from C.elegans nuclei when they were prepared using the X-protocol buffers, but that the higher concentrations of KC1and MgCl in the AP- and A-protocol buffers prevented this loss (e.g., see Paine et al., 1983). Data that is atleast consistent with this hypothesis is the very different KC1 optima of the two types of putative ceHSBFactivity for binding to the HSE probe (Fig. 11).864. The putative HSF activity in the helminth Schistosoma mansoni.The life cycle of S. mansoni is very complex, involving parasitism of molluscan and vertebrate hosts, aswell a nonparasitic, aquatic, larval stage. The regulation of the expression of HSPs in this species is alsocomplex. High levels of S. mansoni heat-shock mRNA are expressed during the cercaria-schistosomuladevelopmental transition, and in adult worms exposed to a 42 °C heat shock (Neumann et al., 1993).EMSA analysis has shown that a putative smHSF activity is detectable in schistosome extracts preparedfrom adult worms kept at 37 °C or 42 °C (control or heat-shock conditions, respectively) (Levy-Holtzman &Schechter, 1994). The protein-DNA complex formed by this putative smHSF activity is also noteworthy fordisplaying an Rf of —0.4-0.5, very similar to that of the protein-DNA complex formed by the putative ceHSBFmactivity.C. THE UTILITY OF KHGlu-BASED ELECTROPHORESIS BUFFERS IN EMSAs.1. EMSA electrophoresis buffers presently in use.Many different electrophoresis buffers have been formulated for use with EMSAs (see Fried, 1989 andLane et al., 1992 for reviews on EMSAs). One of the first of these buffers was TE pH 7.4: 10 mM TRIS, 1 mMNa2EDTA (Fried & Crothers, 1981). The three main types of EMSA electrophoresis buffers presently in use are(i) 0.25-1.Ox TBE buffers (lx TBE is 0.09 M TRIS, 0.09 M boric acid, and 0.0025 M Na2EDTA, pH 8.3;Sambrook et al., 1989); (ii) buffers similar to the one formulated by Strauss & Varshavsky (1984): 6.7 mMTRIS, 3.3 mM NaOAc, 1 mMNa2EDTA, pH 7.5; and (iii) buffers similar to the one formulated by Schneideret al. (1986): 0.38 M TRIS-glycine pH 8.8 in the gel and 40 mM TRIS-glycine pH 8.5 in the buffer chambers(see also Staudt et al., 1986). Of these three types of EMSA electrophoresis buffers the first two are the mostcommonly used.TBE buffers frequently work very well and are the default choice for most investigators. However, theinteraction of borate ions and proteins can give rise to artifactual bands during electrophoresis (Cann, 1966;Parker & Beam, 1963). Borate ion forms covalent complexes with gem-diols, functional groups that are foundon mono- and oligo-saccharides, polyols, and other important biological compounds (Zittle, 1951). Many87eukaryotic transcription factors and nuclear proteins are glycosylated (Jackson & Tjian, 1988, 1989; Kan &cia Silva, 1986; Lichtsteiner & Schibler, 1989), including the only RNA polymerase II regulatory factor fromC. elegans that has been studied biochemically (Okai, 1986a). The use of TBE as an EMSA electrophoresisbuffer may not cause any difficulties when glycosylated sequence-specific DNA-binding proteins are studied,but the potential for difficulties should at least be recognized.The very low ionic-strength EMSA electrophoresis buffers generally lack sufficient pH-bufferingcapacity, and in these cases buffer recirculation is required. However, these buffers are very useful, especially iflow ionic-strength conditions stabilize the sequence-specific protein-DNA interaction under study. Lowionic-strength conditions also minimize the “dead time for the electrophoresis of free DNA into the gel afterapplication of the sample to the well. This “dead time is inversely related to the precision with which certainthermodynamic and kinetic parameters can be measured (Fried & Crothers, 1981).Higher ionic-strength electrophoresis buffers increase the total current resulting from a given appliedvoltage, and reduce the fraction of the current that is carried by charged sample components. These effectsincrease the time required for electrophoresis, as the applied voltage must be reduced in order to preventexcessive heating of the gel. This increased time required for the separation is more than simply inconvenient; itis also viewed as being potentially incompatible with the EMSA analysis of kinetically labile protein-DNAcomplexes. However, the most remarkable aspect of the EMSA technique—and the phenomenon upon which itis based—is the ability of the gel to stabilize the protein-DNA complex throughout the electrophoresis run, evenwhen the half-life of the complex is a very, very small fraction of the time required for the electrophoreticseparation.2. The rationale for the formulation of KHGIu-based EMSA electrophoresis buffers.It is well-known that the ionic milieu can have a very significant influence on protein-DNA interactions(Lohman, 1986; Murdoch et al., 1991; Record et al., 1991; Schroth et al., 1991; Walker & Sikorska, 1986).Consider a purtfled sequence-specific DNA-binding protein interacting with its cognate DNA-binding site. Ifthe salt concentration in the binding reaction is reduced, say, from 150 to 50 mM K+, the binding affinity of theprotein for its cognate DNA-binding site will probably increase (Record et al., 1991). However, at the same88time the binding affinity of the protein for nonspecific DNA may increase to a much greater extent.Sequence-spefic protein-DNA complex formation is therefore not necessarily favoured by lower saltconcentrations, as the determinant of specificity is the ratio of the specific to nonspecific DNA-binding affinities(e.g., see Murdoch et al., 1991). Differential salt effects on nonspecific and sequence-specific DNA-bindingaffinities may be especially significant when studying sequence-specific DNA-binding proteins present in cellextracts, as the total, nonspecific DNA-binding activity present in such extracts vastly exceeds the amount ofsequence-specific DNA-binding activity.The distinctive electrolyte in the EMSA electrophoresis buffers formulated for this thesis is themonopotassium salt of L-glutamic acid (KHG1u). KHG1u provided both a physiological concentration ofpotassium ions (100-150 mM) and a counterion species (glutamate) that is well-known for its ability to stabilizeprotein-DNA interactions (Griep & McHenry, 1989; Guo & Cole, 1989b; Ha et al., 1992; Leirmo et al., 1987;McClelland et al., 1988; Murdoch et al., 1991; Richey et al., 1987; Zou & Richardson, 1991). The latterproperty is important because the EMSA gel must be run for a longer period of time at a low applied voltage(2.5 V/cm).Hydrogen-ion buffering in each of the KHG1u-based buffers was provided by two zwitterionicbuffers—a weak base and a weak acid—whose pKa values were closely matched (see McLellan, 1982). Both ofthese solutes were present at a concentration of 50 mM (20 mM in KGBH) in order to provide a stronglybuffered solution, and buffer recirculation was not required. After 6 h at 40 V the change from the initial pH ineach compartment of the electrophoresis apparatus was ±0.4 pH units (for KGBM or KGTH, using 800 and650 mL of buffer in the lower and upper buffer reservoirs, respectively). KGBM, with an initial pH of —6.3-6.4,was especially useful for the EMSA analysis of the putative ceHSBF-HSE protein-DNA complexes. However,the pH optimum for the analysis of protein-DNA complexes can be quite variable (Carey, 1988; Guo & Cole,1989a; Lane et al., 1992) and should be investigated carefully to avoid artifacts.The KHG1u-based electrophoresis buffers developed for this work do provide a physiologicalconcentration of potassium ions (100-150 mM), but it may be argued that the counterion glutamate isinappropriate for use in studies of sequence-specific DNA-binding proteins from eukaryotic cells. The basis of89this criticism is the fact that although glutamate is a significant intracellular anion in eubacteria (whereaschloride is insignificant), chloride is the major intracellular anion in eukaryotic cells, whereas glutamate isinsignificant (see the discussion in Guo & Cole, 1989, and the references cited therein).Murdoch et aL, (1991) have studied the binding of the X. laevis estrogen receptor (ER) to wild-type(Vit-ERE) and mutant (Mut-ERE) copies of the estrogen response element (ERE) of the Xenopus vitellogeninA2 gene. If a physiological concentration of potassium ion (150 mM) was maintained, the partial replacement ofKC1 by KHG1u had little effect on the relative binding affinity (RBA) of the estrogen receptor for the estrogenresponse element (Table 3). However, if the concentration of potassium ion was reduced from 150 to 50 mM,the RBA of this eukaryotic, sequence-specific protein-DNA interaction dropped precipitously. From these andother data of Murdoch et al. (1991) one may conclude that glutamate is not necessarily inappropriate for use instudies of sequence-specific DNA-binding proteins from eukaryotic cells.Table 3. K}lGlu and the binding of the Xenopus estrogen receptor to the estrogen response element (ERE).KdSalt conditions Vit-ERE (pM) Mut-ERE (nM) RBA50 mM NaC1 580± 85 60± 14 10015OmMNaC1 700± 182 800±460 110050 mM KC1 700± 86 50± 12 7015OmMKC1 320± 63 900± 138 2800100 mMKHG1u + 5OmMKC1 360± 77 900± 170 2500The entire contents of Table 1 of Murdoch et al. (1991) are reproduced above.Vit-ERE: Wild-type ERE of the Xenopus vitellogenin A2 gene.Mut-ERE: Mutant ERE of the Xenopus vitellogenin A2 gene.Kd: Dissociation constant.RBA: Relative binding affinity = {Kd of the Mut-ERE} ÷ {Kd of the Vit-ERE}Finally, it should also be noted that the reverse argument is something that biochemists routinelyencounter whenever they use Type II restriction endonucleases from commercial suppliers. These enzymes are90sequence-specific DNA-binding proteins isolated from eubacterial sources, and hence each of these enzymesfunctions in vivo in an ionic milieu where glutamate—but not chloride—is a prominent anion. Yet in manycases vendors of these enzymes supply their products with a recommended digestion buffer that containschloride ion, because in many cases chloride is quite satisfactory as a replacement for glutamate in reactionbuffers as long as the appropriate cation is present at the appropriate concentration.It is unfortunate that the KHG1u-based buffers used in this thesis were not directly compared with theEMSA electrophoresis buffers currently in use. Ideally such experiments would involve “side-by-side” orparallel EMSA analyses using (i) identical electrophoresis equipment but with running conditions appropriatefor either of the two different buffer systems being compared, (ii) gels prepared using the same polymerizationconditions (%T, %CBjs, and catalyst concentrations) but using either of the two buffer systems being compared,and (iii) loading of the two gels using identical aliquots taken from a series of EMSA binding reactions, each ofwhich would involve a different sequence-specific extract-probe interaction (e.g., using the appropriate probes,and extracts from different sources known to contain the protein of interest). Such parallel EMSA analysescould be carried out, say, using (a) KGBM and TBE, (b) KGBM and the very low ionic-strength bufferformulated by Strauss & Varshavsky (1984), and (c) KGBM and the glycine-containing buffer formulated bySchneider et al. (1986). For comparison purposes it would be essential to compare results obtained usingidentical aliquots from the same EMSA binding reaction. For completeness it would be desirable to set up aseries of distinct binding reactions, so that the various lanes on each gel could be used to determine the relativemerits of the two buffer systems for the study of different sequence-specific extract-probe interactions.The EMSA electrophoresis buffers formulated for this thesis have several disadvantages. They are lessconvenient and more costly to use than lower ionic-strength buffers. They may not be useful for investigationswhich require a minimum dead-thne. The ability of glutamate to stabilize protein-DNA interactions may insome cases be too much of a good thing, as nonspecific protein-DNA interactions are also stabilized by thissolute (Ha et at., 1992). KHG1u-based EMSA electrophoresis buffers are no panacea, but their ability to supplya potassium ion concentration in the 100-150 mM range make them a physiologically relevant option worthconsidering.92CHAPTER 2.1. INTRODUCTIONA. CYTOSINE-SPECIFIC TRANSAMINATION REACTIONS.1. Reactivity of the pyrimidine ring in cytosine derivatives.The reactivity of the pyrimidine ring in cytosine derivatives is influenced by (i) the electron-withdrawing character of the 1 ,3-diazine ring nitrogens, which renders the carbon atoms at the 2, 4, and 6positions particularly electron-deficient; (ii) the participation of the 5,6 double bond in a conjugated lactam,which increases the electron deficiency at C-6; (lii) the participation of the exocyclic nitrogen (N-4) in aconjugated system with the carbonyl at C-2, which significantly increases the (otherwise negligible) ability ofthe former to act as a leaving group; and (iv) the susceptibility of N-3 to protonation and alkylation (Shepherd &Fedrick, 1965; Ueda, 1988; Vorbruggen, 1990). An important consequence of the last point is that under mildlyacidic conditions it is N-3 (pKa = 4.5 in cytidine), not N-4, which becomes protonated, although the resultingpositive charge can be delocalized to N-i, N-3, and N-4 (Ueda, 1988). Hence protonation of N-3, saturation ofthe 5,6 double bond, or addition across the 5,6 double bond can all accentuate the electron deficiency of thecytosine ring and facilitate nucleophilic substitution at C-4.2. The Bucherer reaction.The Bucherer reaction can be viewed as the reversible, bisulfite-catalyzed interconversion of aromaticamities and hydroxyaromatic compounds (Seeboth, 1967). A compound such as napthol can be used to prepareprimary, secondary, and tertiary napthylamines; conversely, napthylamine can be deaminated or converted tosecondary or tertiary napthylamines (see Fig. 16). A remarkable feature of these reactions is that they proceedvia tetralonesulfonic and tetraloniminesulfonic acid intermediates, and not via bisulfite addition to the keto formof napthol or the imine form of napthylamine.93+ HS03- HS03DeaminationNH2OHOH- HS03 —— so3- + HS031 L H2N OHcJtx% - NH3 SO3-+H2Oj -H2ONH2cxLTransamination- NH3NH+RNH2 -RNH2H2N NHRSO3-NHR+NH3- HSO3so3- ÷HSOFigure 16. The Bucherer reaction. Adapted from Seeboth (1967).NH— HNc÷H20B ONB -H20+RNH2I[.RNH2TransaminationNRHN”11N -NH3ON”B +NH3H2N NHRO’BN4-Alkylcytosinederivative94DeaminationNH2N0- HBHN”+HB 0N’”B!NH3Uracil derivativeNH2+ HB- HBH2N OHO’BNHR NHR- HBFigure 17. The AE mechanism for the deamination and transanuination of cytosme derivatives.Adapted from Garrett & Tsau (1972), Shapiro & Klein (1966), and Shapiro & Weisgras (1970).953. The deamination and transamination reactions of cytosine derivatives.There are important parallels between the Bucherer reaction and the reaction of cytosine derivativeswith bisulfite (Hayatsu et al., 1970; Notari, 1967; Shapiro & Weisgras, 1970; Shapiro et al., 1970). Bothreactions proceed via an addition-elimination (AE) mechanism (Hayatsu, 1976a; Seeboth, 1967; Shepherd &Fedrick, 1965; Slae & Shapiro, 1978; Vorbruggen, 1990). The initial step of both reactions is the reversibleaddition of bisulfite at a position meta to a carbon atom bearing a heteroatom conjugated to the ring. Saturationof the 5,6 double bond of cytosine, and in particular the addition of bisulfite across this bond, dramatically shiftsthe tautomeric equilibrium of cytosine in favour of the imine form (see Table 4). Like napthylamine, cytosinemay then be deaminated (to uracil) or transaminated (to anN4-alkylcytosine derivative), depending on thereaction conditions (see Fig. 17). The final step of both reactions is the base-catalyzed elimination of bisulfite toform the product.The choice of reaction pH for a bisulfite-catalyzed, cytosine transamination reaction has an importantinfluence on the ratio of deaminated to transaminated products (Hayatsu, 1976a). Transamination reactions aretypically performed at pH 6.0 to 7.5. The pH optimum of a transamination reaction presumably reflects threephenomena: the decrease in the equilibrium constant for formation of the 5,6-dihydrocytosine-6-sulfonateintermediate with increasing pH, and protonation of the transaminating amine (to form the unreactive conjugateacid) and sulfite ion (pIta 6.24, which catalyzes the hydrolysis of the 5,6-dthydrocytosine-6-sulfonateintermediate) with decreasing pH (Hayatsu, 1976a).An important feature of bisulfite-catalyzed deamination and transamination reactions is that they arevery specific for the cytosine ring if they are carried out in the presence of an antioxidant (Hayatsu, 1976a).Under these conditions other heterocycles and nonheterocyclic residues found in nucleic acids do not react(although bisulfite will add reversibly across the 5,6 double bond of uracil). The presence of an antioxidant isrequired because the autoxidation of bisulfite ion releases free radicals that degrade polynucleotides and theirconstituent monomers (Hayatsu, 1976a).Bisulfite is not the only compound whose addition across the 5,6 double bond catalyzes thedeamination and transamination of cytosine derivatives. A variety of buffers were found to act as weak catalysts96for the deamination of cytosine at pH<6.O, conditions which would result in the partial protonation of N-3(Garrett & Tsau, 1972; Shapiro & Klein, 1966). However, the most effective catalysts from among these bufferswere mercapto acids, e.g., mercaptoacetic acid (Shapiro & Klein, 1966). It was later found that manysulfhydryl-containing small molecules add across the 5,6 double bond and catalyze the deamination andiransamination of cytosine derivatives (Boni & Budowsky, 1973; Hayatsu, 1976a; Wataya et al., 1973).Table 4. Tautomeric equilibria of cytosine derivatives m aqueous solution.Pyrimidine moiety Enamme tautomer Imine tautomer Enamine to imine ratio5,6-Dthydrocytosine-6-sulfonate, pH 5.1NH2N’NH2H?NrLsoa÷NH2so3-<1(estimated)NH2 NHCytosineN4-Aminocytosine5,6-DihydrocytosineNH2HN’to 103025NHData are from Brown et a!. (1968), Kwiatkowski & Pullman (1975) and Taguchi & Wang (1977).97The active-site cysteine sulfhydryl groups of a variety of enzymes are also believed to add across the5,6 double bond of pyrimidine derivatives during the course of their normal enzymatic functions in vivo.Members of this class of enzymes include thymidylate synthase (Hardy et at., 1987; Kawai & Hilicoat, 1974),DNA (cytosine-5) methyltransferases (Chen et at., 1991; Friedman & Ansari, 1992; Hanck et at., 1993;Klimasauskas et al., 1994; Kumar et at., 1994; Smith et at., 1992; Wyszynski et a!., 1992, 1993), bacterialcytidine cleaminases (Yang et at., 1992), and eukaryotic cytidine deaminases that act on specific cytidylateresidues in certain mRNA transcripts (e.g., see Hodges & Scott, 1992).The bisulfite-catalyzed deamination of 5-methylcytosine to thymine is very inefficient, primarilybecause of a reduction in the rate of formation of the 5,6-dihydrocytosine-6-sulfonate intermediate (Wang et at.,1980). This difference in the reactivity of cytosine and 5-methylcytosine has been used to map5-methylcytosine residues in genomic DNA. In this procedure genomic DNA is denatured, cytosine residues arequantitatively deaminated to uracil, and the resulting DNA is amplified by the polymerase chain reaction to yielda positive display of “C” residues that actually represent 5-methylcytosine residues that were unaffected by thedeamination step (Frommer et at., 1992).Several other reports have described practical applications of the bisulfite-catalyzed transamination ofcytosine derivatives. Ethylenediamine and similar diamines have been used to Iransaminate cytosine derivativesand allow for the subsequent attachment of reporter groups. Draper (1984) introduced a fluorescent dye intospecific positions of an RNA transcript using this method, and Miller & Cushman (1992) later reported avariation of this derivatization procedure using oligonucleotides. Biotin N-hydroxysuccinimide ester has beenlinked to cytosine residues in DNA (Viscidi et at., 1986) and cytosine-containing nucleotides (Gillam & Tener,1986) using similar derivatization procedures.4. The transamination of cytosine derivatives by acyl hydrazides.a. The chemistry ofacyl hydrazides.Smith (1983) has succinctly described acyl hydrazides as “acyt hydrazines, compounds derivedfrom anacid byformal replacement of a leydroxyt group by a hydrazino group, which may itseifbear other substituents”.98Some noteworthy structural features of a simple carboxylic acid hydrazide (acethydrazide) are summarized inFig. 18. The nitrogens and the carbonyl group are essentially coplanar, and the hydrogens on the terminal (orN’) nitrogen are pointed towards the oxygen atom (Paulsen & Stoye, 1970). The lengths of the C-N and N-Nbonds indicate that both possess some double-bond character (Smith, 1983). The pKa of the conjugate acid ofacethydrazide is 3.24, which reflects the weakly basic character of most simple monoacyl hydrazides (Lindegren& Niemann, 1949).:0:4,.,HC—C\ \. Locants::N—N: pKa3.24I \ (conjugate acid) N N’ 1IUPAC) 1’ 2’ J1.33 A 1.40 A 1 2 ChemicalAbstractsFigure Th. Structural features of acetic acid hydrazide (acethydrazide).The six atoms shown in boldface lie almost exactly in the same plane. In the absence of functional groups ofhigher priority, the locants N and N’ refer to the acylated and the terminal nitrogens, respectively. The bondlengths shown will vary slightly depending on the hydrazide, but in all cases these lengths are shorter than thecorresponding single-bond lengths. Adapted from Paulsen & Stoye (1970) and Smith (1983).Paulsen & Stoye (1970) and Smith (1983) have written excellent reviews of the reactivity of acylhydrazides. Acyl hydrazides are stable to heating (below 180 °C), and acid- or base-catalyzed hydrolysis isslow, especially with N’-substituted derivatives. A more commonly observed decomposition reaction isoxidation. The N’ nitrogen is nucleophilic, but it does not react readily with alkylating agents. Unsubstituted ormonosubstituted acyl hydrazides can be acylated by acid anhydrides and acid chlorides. Unsubstituted acylhydrazides also react with nitrous acid to form acid azides. However, the best-known reaction of unsubstitutedacyl hydrazides is the formation of acyl hydrazones from aldehydes and ketones. The equilibrium constants formost acyl hydrazone reactions strongly favour product formation.99(a) (b) (c)Figure 19. The reaction of Girards reagent P with cytosine derivatives.Adapted from Kikugawa et aL (1967a).b. The reaction ofacyl hydrazides with cylosine derivatives.Girard’s reagents are acyl hydrazides that contain a quaternary ammonium substituent (Girard &Sandulesco, 1936). These compounds are usually used to convert aldehydes and ketones to water-soluble acylHHGirard’s reagent PHHN”RpH 4.2Where R = a, b, or C:IO=I,—O.0H( HiHHO OHI_O—P— OII00100hydrazone derivatives for isolation purposes. Girard’s reagent P (acethydrazidopyridinium chloride) was laterfound to act as a transaminating reagent for a variety of cytosine derivatives (Kikugawa et al., 1967a, 1967b).This reaction (see Fig. 19) has a very sharp pH opthnum (4.2), which presumably reflects two phenomena:protonation of the cytosine ring under mildly acidic conditions (recall that the pKa of N-3 in cytidine is 4.5),which should increase the reaction rate, and protonation of the acyl hydrazide under increasingly acidicconditions (recall that the pKa of acethydrazide is 3.24), which should decrease the reaction rate (Gal-Or et al.,1967; Kikugawa et a!., 1967a).Malonyl hydrazide and acethydrazide also transaminate cytosine derivatives under mildly acidicconditions (Gal-Or et a!., 1967). Semicarbazide (carbamohydrazide,H2NCONHNH)is also an effectivetransaminating reagent for this reaction (Hayatsu et a!., 1966; Hayatsu & Ukita, 1964, 1966). As expected,bisulfite can act as a catalyst for the reaction of semicarbazide with cytosine derivatives (Hayatsu, 1976b).Bisulfite has also been used to catalyze the reaction of biotin hydrazide with cytosine residues inoligonucleotides (Reisfeld et a!., 1987); however, bisulfite catalysis is not an essential requirement for thereaction (Gal-Or et al., 1967; Hayatsu, 1976a, 1976b; Kikugawa et a!., 1967a, 1967b).An important feature of this reaction is that it is very specific for the cytosine ring. Other heterocyclesand nonheterocyclic residues found in nucleic acids do not react with acyl hydrazides, even after prolongedincubation (Gal-Or et a!., 1967; Hayatsu et a!., 1966, 1976b; Hayatsu & Ukita, 1964, 1966; Kikugawa et a!.,1967a, 1967b; Reisfeld et a!., 1987). Another noteworthy feature is the relative inefficiency of the competingdeamination reaction (Gal-Or et a!., 1967; Hayatsu et al., 1966; Hayatsu & Ukita, 1964).5. Reactivity of cytosine moieties in polynucleotides.There are many common features shared by the two types of cytosine-specific transamination reactionsdescribed above (Hayatsu, 1976a, 1976b). Their reaction mechanisms are believed to be similar, and bothreactions are very specific for cytosine residues in nucleic acids. Another common feature is the lack ofreactivity of cytosine residues in double-stranded polynucleotides. The involvement of cytosine in Watson-Crick base pairing causes the pKa of N-3 to drop from approximately 4.6 in single-stranded DNA to 3.7 indouble-stranded DNA (Banville et a!., 1986; Rawitscher & Sturtevant, 1964). The hydrogen bonding101interactions of N-3 and N-4 of cytosine moieties with the N-i hydrogen and 0-6 of guanine moieties,respectively, are preserved under the reaction conditions employed by either type of transamination reaction.Thus in the absence of helix-coil transitions induced by DNA supercoiling, elevated temperatures, or highconcentrations of helix-destabilizing solutes there is no detectable reactivity of cytosine residues indouble-stranded regions of polynucleotides (Draper, 1984; Hayatsu, 1976a, 1976b; Hayatsu & Ukita, 1966;Shapiro et al., 1973).B. CYANATE ESTERS, ACYL HYDRAZIDES, AND THE IMMOBILIZATION OF NUCLEIC ACIDS ON SOLIDSUPPORTS.1. Strategies for the immobilization of DNA on chromatography resins.Various methods have been developed for the immobilization of DNA on hybridization membranes andrelated solid supports. However, these immobilization methods will not be considered in this discussion.Instead I will focus on those methods that have been used for the immobilization of DNA for the affinitychromatography of DNA-binding proteins.Currently there are three major strategies for the immobilization of double-stranded DNA on solidsupports for the affinity chromatography of DNA-binding proteins. One strategy is the introduction offunctional groups on the resin, such as cyanate esters (Arndt-Jovin et al., 1975; Poonian et al., 1971) or epoxides(Inomata et al., 1992), that are able to react with residues in unmodified DNA. Another strategy is to incorporatebiotinylated deoxyribonucleotides into synthetic or natural DNA, and then immobilize the modified DNA usingavidin- or streptavidin-derivatized solid supports (e.g., Kasher et al., 1986). A third strategy is to incorporate atethered, chemically reactive functional group (e.g., a primary amine; Larson & Verdine, 1992) into syntheticDNA, and then immobilize the modified DNA using an appropriately derivatized solid support.2. The use of CNBr-activated agarose for the immobilization of unmodified DNA.The overwhelming majority of recent studies on the affinity chromatography of sequence-specificDNA-binding proteins have used CNBr-activated Sepharose (Sepharose is the brand name of the most102commonly used beaded agarose resin) (Kadonaga, 1991; Kadonaga & Tjian, 1986). In these experimentsSepharose is freshly activated with —100 mg of CNBr per millilitre of washed resin and then used immediately(Kadonaga, 1991). This is an extremely high degree of activation, as commercially activated Sepharose 4Bresins are prepared using only 5-10 mg of CNBr per gram of drained resin (Wilchek et at., 1984). About60-85% of the coupling capacity of freshly activated Sepharose is attributable to cyanate esters, with theremainder attributable to much less reactive imidocarbonates (Wilchek et at., 1984). These cyanate esterspresumably react with an exocycic amine present on cytosine, guanine, and especially adenine residues in theDNA (Poonian et at., 1971), and if such a reaction occurs an isourea linkage is formed (Wilchek et at., 1984).3. The use of acyl hydrazide-derivatized agarose for the immobilization of oxidized RNA.The utility of acyl hydrazide-derivatized solid supports for the affinity chromatography ofRNA-binding proteins has already been demonstrated. Lamed et at. (1973) took the resourceful step ofconverting a commercially available, CNBr-activated Sepharose matrix into an acyl hydrazide resin using adipicacid dihydrazide (which also acted as a “spacer arm), and resins prepared using this method have proven usefulfor various affinity chromatography applications (Burrell & Horowitz, 1975, 1977; Ulbrich & Wool, 1978;Ulbrich et at., 1979). In the above reports and related studies (Robberson & Davidson, 1972; Vasilenko et at.,1972), ribonucleotides or purified RNA preparations were treated with periodate (a mild oxidizing agent that isrelatively specific for cis-diols) to yield mono- or poly-nucleotides with reactive, 3’-terminal dialdehydes. Thesemodified RNA molecules were immobilized on acyl hydrazide-derivatized solid supports via the formation ofstable acyl hydrazone bonds. This type of covalent linkage has also been used to immobilize mildly oxidizedglycoproteins to acyl hydrazide-derivatized solid supports (reviewed in O’Shannessy & Wilchek, 1990), but thevery nature of DNA precludes its immobilization via this type of linkage.Acyl azides (RCON3)are formed when an acyl hydrazide-derivatized resin is treated with nitrous acid(Suttnar et at., 1977). Although acyl azide derivatized resins have been synthesized as intermediates (i.e.,subject to further derivatization) and subsequently used for the immobilization of ligands for affinitychromatography, these resins have never been used for the immobilization of DNA.103C. SUMMARY OF THESIS OBJECTIVES FOR PART 2.The only commonly used chemistry for the covalent immobilization of unmodified, double-strandedDNA on solid supports is CNBr activation of beaded agarose. I wanted to investigate the potential of acylhydrazide-mediated transamination reactions for this purpose. Although the objective of these experiments wasto covalently immobilize sterically accessible, double-stranded DNA for the affinity chromatography ofsequence-specific DNA-binding proteins, I also tried to illustrate how DNA immobilized using this reactionchemistry might be a suitable substrate for analytical methods in other important areas of biomedical research.104CHAPTER 2.2. MATERIALS AND METHODSA. MATERIALS AND ROUTINE EXPERIMENTAL PROCEDURES.These are described in Part 1 of this thesis.B. THE IMMOBILIZATION OF DNA ONACYL HYDRAZIDE-DERIVATIZED SOLID SUPPORTS.1. Preparation of DNA for use in immobilization assays.a. Very large-scale preparation ofpUC 19 plasmid DNA.Highly purified pUC 19 plasmid DNA was prepared from 3.2-litre “fermentor” culturest of E. coliDH5a::pUC 19 grown at 37 °C for 22 hours in ‘Terrific Broth” (Tartof & Hobbs, 1988) plus ampicillin(0.050 mglmL). Described below is a very large-scale, modified version of the well-known alkaline-lysisprocedure for purifying plasmid DNA (Birnboim & Doly, 1979; Sambrook et al., 1989). Typical yields were20-40 mg of very high purity plasmid DNA per 3.2-litre culture.Chilled bacteria were centrifuged using four (tared) 250-niL bottles (3 spins at 5,000 rpm, 15 mm, 4 °C,using a Sorvall GSA rotor and RC-5 centrifuge) and then washed once with 200 niL of ice-cold STE pH 8.0 perbottle. Bacterial pellets (-18 g each) were softened by vortexing, and then 50 mL of ice-cold, alkaline-lysisSolution I (with freshly added lysozyme, 4 mg/mL) was added per bottle and mixed immediately. After 6 mm at20-22 °C, 100 mL of freshly prepared Solution II was added per bottle and mixed immediately by inversion(20x), and then the bottles were placed on ice. After 10 mm, 75 mL of Solution III was added per bottle andmixed immediately by inversion (50x), and then the bottles were placed on ice again. After 10 mm, debris waspelleted for 90 mm at 13,000 rpm at 4 °C in a GSA rotor. The supernate (-900 mL) was transferred to anddivided equally among six 250-mL bottles, and then 90 mL of 2-propanol was added per bottle and mixed well.After 30 mm at 20-22 °C, nucleic acids and residual protein were pelleted for 30 mm at 9,000 rpm at 20-22 °Ct Improvised equipment was used as a matter of necessity. The same culture vessels used for the large-scalegrowth of C. elegans were used here.105in a GSA rotor. All traces of the supernate were discarded, and then50 mL of TE pH 7.4 was added toresuspend each of the pellets at 4 °Covernight.The next day 25 mL of 7.5 M NH4OAc was added per bottle andmixed, and then the bottles wereplaced on ice for an hour. High-Mr RNA and protein were then pelleted for 30 mm at 9,000 rpm at 4 °C in aGSA rotor. The supernate (-440 mL) wastransferred to and divided equally among four 250-mL bottles, andthen 1 vol of 2-propanol was added per bottle and mixed. After 30 miii at 20-22 °C, DNA was pelleted for 30miii at 9,000 rpm at 4 °C in a GSA rotor. All traces of the supernate were removed, and each pellet wasresuspended with 18.3 niL of TB pH 8.0. Resuspended DNA was pooled in one bottle, and 76 g of CsC1 wasadded. After the CsC1 was dissolved, the solution was divided equallybetween two 250-niL bottles, and EtdBr(4.0 mL of a 10 mg/mL stock) was added to each bottle and mixed. After 30 miii in the dark at 20-22 °C,RNA-EtdBr complexes were pelleted for 30 mm at 9,000 rpm at 20-22 °C in a GSA rotor. The supernatewascarefully transferred to a 250-niL, vacuum-driven sterile-filtration unit (Corning), and protein-EtdBr complexeswere removed by filtration undersubdued light. High purity Triton X-l00 (110 iL of a 10% w/v stock) wasadded to the filtrate, the solution was mixed, and the density of the solution was confirmed to be, or adjusted to,1.54-1.56 gImL. The DNA was thencentrifuged to equilibrium in eight 13.5-mL Quick-Seal tubes (Beckman)for 24 hours at 60,000 rpm at 20 °Cusing a Beckman Ti 70.1 rotor and L8-70 centrifuge.The lower, supercoiled-DNA-contauning band was removed from each tube, the DNA-rich fractionswere pooled, and then the EtdBr wasextracted seven times with 1-butanol.These procedures were performedunder subdued light. The DNA wasdiluted to 90 mL with TB pH 8.0, transferred to a 250-niL bottle, and thenprecipitated with 0.1 vol of 3.0 M NaOAc pH 5.2, and one volume of 2-propanol. The DNA was pelleted fromthe 250-mL bottle, and later reprecipitated from a 40-niL Oakridge tube using NaOAc and EtOH.b. Restriction endonuclease digests ofDNA.Plasmid DNA, in amounts of 12.0 or 16.0 mg, was typically digested with15,000 or 20,000 U of Xba Ior Kpn I, respectively (i.e., —1.2 U of enzyme per .tg of DNA) for 8 hours in a 37 °C water bath. Reactions(24.0-mL final volumes in 40-niL tubes) were carried out in lx reaction buffer and0.10 mg/niL of acetylated(nuclease-free) BSA (supplied as lOx and lOOx stock solutions by the vendor, respectively).106Restriction endonuclease activity was stopped by the addition of 0.60 mL of 500 mM Na2EDTA pH8.0. Any traces of residual, 10W-Mr RNA were digested by adding ribonuclease A (0.40 mL of a 1.0 mglmLstock) and continuing the reactions a further two hours at 37 °C in a water bath. Aliquots (1.5 or 2 liL) fromeach reaction were analyzed by agarose gel electrophoresis to confirm that the restriction endonuclease digestswent to completion. Fifteen miliitres of 2.5 M NaC1, 20% wlv PEG 8000 was added per reaction and then theDNA was precipitated overnight at 4 °C.The DNA was pelleted and all traces of the supernate were removed. Each DNA pellet wasresuspended with 12 mL of TE pH 8.0, and residual protein was extracted at least three times withphenol:chloroform:isoamyl alcohol (25:24:1, vlvlv). The DNA was then precipitated, pelleted, and all traces ofthe supernate were removed. Each DNA pellet was rinsed thoroughly with 40 ml. of 70% EtOH and thenrepelleted. All traces of the supernate were removed, and the pellets were dried 15-20 mm. The DNA pelletswere resuspended with 10 mM KC1 to a concentration of —2.1 to 2.5 mglmL (as determined by UV-ABSspectrophotometry). Each preparation was assigned a lot number (e.g., K1, Xl, etc.), and each lot was dividedinto fixed-volume aliquots (typically 0.36-mL) and stored at -20 °C.Small-scale digestions of pUC 19 DNA were performed with a variety of restriction endonucleases (seeTable 12). These digests were carried out with 1,000 U of enzyme (2,500 U for the Sal I digest) and 0.50 mg ofDNA. Reactions (2.4-mL final volumes in 14-mL tubes) were carried out for 16 hours at the recommendedtemperature in lx reaction buffer and 0.10 mg/mL of acetylated (nuclease-free) BSA (supplied as lOx and lOOxstock solutions by the vendor, respectively). Stopping of restriction endonuclease activity, nbonuclease Atreatment, analytical gel electrophoresis and PEGINaC1 precipitation were performed essentially as describedabove, except that the reagent volumes added were reduced tenfold. The DNA (3x 0.83 mL) was precipitatedwith PEG/NaC1 (3x 0.50 mL) in three microcentrifuge tubes. After centrifugation and removal of the supernate,each of the three DNA pellets was resuspended with 400 IlL of TE pH 8.0. The DNA solutions derived from agiven digest type were pooled, and residual protein was extracted at least three times withphenol:chloroform:isoamyl alcohol (25:24:1, vlvlv).107The DNA was then precipitated, pelleted, and all traces of the supernate were removed. Each DNApellet was rinsed three times with 1.0 mL of 70% EtOH and repelleted. All traces of the supernate wereremoved, and the pellets were dried 10 mm. The DNA pellets derived from a given digest type wereresuspended with 10 mM KC1, and then the resulting solutions were pooled to give a DNA concentration of4.8 mg/mL (as determined by UV-ABS spectrophotometry). The DNA solutions were stored at -20 °C.c. Preparation ofpoly(dI-dC)•poly(dI-dC).Poly(dI-dC)poly(dI-dC) (25 units) was resuspended with 1.9 mL of KBHE and left at 4 °C for severaldays. The DNA was precipitated (in 3x 0.63-mL aliquots), left for 10 mm, and then microcentrifuged 15 mm at20-22 °C. The pellets were washed 3x with 1.5 mL of 70% EtOH, and all traces of the supemate were removedafter the initial pelleting and after each wash. The pellets were dried 10 mm, and then each pellet wasresuspended with 200 jiL of 10 mM KC1. Resuspension of the DNA required heating at 50 °C for 10-20minutes. The DNA was then allowed to cool slowly to 20-22 °C (i.e., the heating block was turned off and leftto cool to room temperature). The resuspended DNA was pooled, left on ice for an hour, and then itsconcentration was determined spectrophotometrically. The DNA solution was stored at -20 °C in 0.20-mLaliquots.2. Handling of small quantities of chromatography resins.This thesis describes the routine manipulation of very small quantities of chromatography resins. Thepipetting of Affi-Prep HZ (rigid beads with diameters of 0.04-0.06 mm) does not usually present any problems,especially when the resin is properly resuspended in buffers containing PEG. Special wide-bore pipette tips(Bio-Rad BR-34, Cat no. 223-9034) were used to dispense small aliquots of agarose-based resins (soft beadswith diameters of 0.075-0.300 mm).Please see Part 1, Chapter 1.2[B.4-51 of this thesis for definitions of the terms “mixing by rotation” and“degassing of solutions or chromatography resins”. The term “the resin was pelleted” denotes centnfugation ina clinical centrifuge at 20-22 °C at about 1700 rpm for 30 seconds (if no other length of time was specified).108Each resin-containing cryotube was placed on top of an empty cryotube inside of a 14-mL tube to allow forconvenient pelleting of the resin.Resins were washed at 20-22 °C by combining the indicated amount of resin (as little as 40 j.iL) withthe indicated amount of buffer (typically 2.0 to 12 mL) in the indicated size of disposable polypropylene tube(typically cryotubes or 14-mL tubes). The resin was gently resuspended (by manually inverting the tube), mixedby rotation for 10 mm, pelleted, and the supernate was removed by aspiration.3. The solubility of TCBA salts at pH 4.2.All 1.25x coupling buffers were adjusted to give a stable reading of pH 4.20 and then filter-sterilized.When TCBA was used as a hydrogen-ion buffer there was a small amount of a very fine precipitate left on thefilter, but the removal of this material did not affect the pH of these buffers. Thus the actual concentration ofTCBA may have been slightly less than the nominal concentration of this solute given in Tables 2 and 4-9.4. The standard DNA-immobilization reaction.This procedure is summarized in Fig. 20 and described in detail below.Affi-Prep HZ and Affi-Gel HZ resins were stored at 4 °C as —50% v/v suspensions in 2-propanol.Small aliquots of these acyl hydrazide-derivatized resins were washed by rotation at 20-22 °C with 1 .25xcoupling buffer (CB) just before use in DNA-immobilization reactions. Various coupling buffers wereexamined for their effect on coupling efficiency, and one set of five reactions was used to examine a singlepreparation of a given coupling buffer. Described below is the resin washing procedure used for each set ofreactions.An aliquot of well-suspended resin suspension (containing —0.3 to 0.6 mL of resin) was washed fourtimes with 12 mL of 1 .25x CB in a 14-mL tube. After the fourth wash, the resin was resuspended with 1 mL of1 .25x CB and then poured into a cryotube that had been tared on an analytical balance. The resin was pelletedand all of the supernate was removed. The wet resin was weighed on an analytical balance, and then 1.00 mL of1.25x CB was added per gram of wet resin. The resin was resuspended, quantitatively transferred and depositedat the bottom of a 50-mL tube, and then aseptically degassed for 5 mm. The degassed resin was gently but109uniformly resuspended prior to the removal of each of five 80-.iL aliquots of suspension, which were transferredto cryotubes. The nominal concentration of degassed resin in these aliquots was 50% v/v.Aliquots (typically 0.36-mL) of linearized DNA (1.8—2.4 mg/mL in 10mM KC1) were thawed, mixed,and then microcentrifuged 2 mm at 20-22 °C. DNA (20 pL per reaction) was added to four of the five 80-IlLaliquots of 50% v/v resin suspension of each set of reactions. A single, mock immobilization reaction wasperformed using the fifth aliquot of resin suspension of each set of reactions (i.e., 20 iiI. of 10 mM KC1 wasadded instead of DNA). Unless specified otherwise, reactions were mixed by rotation for 6 hours in a preheated,37 °C incubator (30 °C for Affi-Gel HZ).DNA-immobilization reactions were stopped with the addition of 2.00 mL of SB (see Table 2). Thestopped reactions were mixed by rotation for one hour as before (at 37 °C; 30 °C for Affi-Gel HZ). The resinwas pelleted for 2 mm, 1.0 mL of the supernate was transferred to a microcentrifuge tube, and then theconcentration of uncoupled DNA in each SB-stopped reaction was determined by UV-ABS spectrophotometry.The total volume of the supernate was taken to be 2.10 mL for calculation puiposes. The spectrophotometer waszeroed with SB, and the absorbance values of the supernates of a given set of quadruplicate reactions (i.e., thatexamined a given coupling buffer) were corrected using the absorbance value of the supernate of the appropriateSB-stopped, mock immobilization reaction.One set of “no-resin control” (NRC) reactions was performed for each lot of DNA used. Thesereactions were performed exactly as described above, except that 80-jiL aliquots of 1 .25x coupling buffer wereused instead of resin suspension. The absorbance values of a given set of quadruplicate NRC reactions werecorrected using the absorbance value of an SB-stopped, mock NRC reaction (i.e., where 10 mM KC1 was addedto the 1.25x coupling buffer instead of DNA). The result obtained from the appropriate set of NRC reactionswas used to estimate the yield and efficiency of all immobilization reactions that used a given lot of DNA.The DNA-inunobilization reactions carried out using lot Ki DNA were performed as described abovewith the following modifications. Lot Ki DNA was degassed for 5 mm prior to use, and a rotation speed of 50rpm was used for washing the resin and for the immobilization (4-16 h) and stopping reactions (2 h).110Immobilization reactions__Ibi idWash the resin Prepare 50% v/v Add 80 juL of the 50% v/v resinwith 1 .25x CB resin suspension suspension to tubes R0 to R4(coupling buffer)No-resin cono1 reactions Add 20 L of 10 mMr4 100-iiLeactionsKC1 to tubes R0 and N0Add2OjuLofDNA bI I inlOmMKCltoAdd 80 iL of 1.25x CB \J tubesR14andNto tubes N0 to N4Idi lel ifiAN={A260 of N1 4) —{A260 of N0)ARfA2600fR1}_{f O}/ 2.10-mLQ stopped 0 ‘26O = (AN — AR)\ reactions jQuantitate the immobilizedAdd 2.00 ml of DNA spectrophotometricallystop buffer per tubeIgi Ihi IiiFigure 20. Protocol for microscale DNA-immobilization reactions.1115. Storage of immobilized DNA and the assay of linkage stability.Standard DNA-immobilization reactions were performed using buffer TC and either linearized pUC 19DNA or poly(dI-dC).poly(dI-dC). Immediately after the initial stop reaction the remaining supernate (-4 mL)above aliquots of resin (40 l.tL, containing quantitated amounts of immobilized DNA) was removed byaspiration and the resin was washed three times with 2.0 mL of SB. After the supernate from the last wash wasremoved, 2.00 mL of SB and 160 pL of 700 mM NaF were added to each tube. Resin aliquots were stored at4 °C shielded from light.The percentage of immobilized DNA released after storage at 4 °C for ten months was assayed toevaluate the stability of the covalent linkage. After ten months at 4 °C the stored resin suspensions were firstequilibrated to 20-22 °C, mixed by rotation, pelleted, and then the DNA was quantitated spectrophotomeiricallyas before. In this assay the total volume of each supemate was taken to be 2.2 mL for calculation purposes.C. IMMOBILIZED DNA AS AN AFFINITY CHROMATOGRAPHYLIGAND.1. Construction of an HSE-affinity resin.Except as noted, all of the buffers and procedures described below were at 20-22 °C.The hse-A.hse-T duplex was first polymerized to form head-to-tail repeats (using 1.0 mg = 30 nmol ofthe 5 1-bp monomer and 70 U of T4 DNA ligase per 1 .0-mL overnight reaction, performed in the presence ofBSA and PEG as described in Sambrook et al., 1989). At the end of the ligation reaction, proteins wereextracted six times with phenol:chloroform:isoamyl alcohol (25:24:1, vlv/v). The DNA was precipitated for 30mm with NaOAc and 2-propanol, and then microcentrifuged for 10 mm, at 20-22 °C. The DNA pellets wereeach washed three times with 1.0 mL of 70% EtOH, dried, and then resuspended to a concentration of—2.0 mg/mL with 10mM KC1.The polymenzed hse-A•hse-T duplex was immobilized on Affi-Gel HZ (lot 43482B; see Chapters2.2[B.4] and 2.3[A.3J of Part 2 of this thesis). The standard immobilization reaction was scaled up 20-fold (to afinal volume of 2.0 mL). The DNA was immobilized on the resin using buffer TC for 6 hours at 30 °C in a50-mL tube. The immobilization reaction was stopped with 40.0 mL of SB for 1 hour at 30 °C, the resin was112pelleted, and then the uncoupled DNA was assayed spectrophotometrically. No attempt was made to “cap”unreacted acyl hydrazide groups. The resin (-0.42mg DNAImL) was washed 3x with 50 mL of SB, 2x with50 mL of CEB-200012% w/v PEG 8000, and then stored in the dark at 4 °C under -3 mL of CEB-200012% wlvPEG 8000.The next dayt the HSE-affinity resin was equilibrated to 20-22 °C, degassed, and then all of thesupernate was removed. A 10-mL, polypropylene chromatography column (Poly-Prep, Bio-Rad) was carefullypacked with the HSE resin (-0.8-mL bed volume). The packed column was washed with 10 ml. of degassedCEB -2000 and then 10 mL of degassed CWB at a flow rate of -0.30 mL/min using a peristaltic pump connectedto the Luer outlet of the column.2. Microscale affinity chromatography.Two -1 .0-mL aliquots of HS nuclear extract (prepared using the A-protocol as described in Chapter1.2[C.3] of Part 1 of this thesis) were thawed for 2 mm at 37 °C and then pooled in a 14-mL tube containing a3x10 mm “flea” stir-bar. The following solutions were added dropwise while the extract was stirred moderately(0.6x max. speed): 0.50 mL of 2.0 mg/mL pdN6,0.050 mL of freshly prepared 20% w/v n-octyl glucoside, andthen water to bring the volume of the solution to 5.0 mL. Stirring was stopped, the stir-bar was removed, andthen the 5-mL solution was distributed among four microcenthfuge tubes. Insoluble proteins were pelleted in amicrocentrifuge at 4 °C for 15 mm. The supernate was carefully transferred to a 20-mL disposablepolypropylene syringe equipped with a filtration device (450-nm pore size, low protein-binding, 13-nun dia.,Millex GV13,Millipore), and the supernate was filtered into a 14-mi. tube.The filtered supernate was very carefully loaded on and run into the HSE-DNA-affmity column. Theaffinity column was run at -0.10 mL/min using a peristaltic pump connected to the Luer outlet of the column.The affinity column was washed with 10 mL of degassed CWB containing 0.2 mg/mL of pdN6, and then with10 mL of degassed CWB. The column was washed using a flow rate of .-0.20 mL/min.t Do NOT store the resin in solutions containing PEG for other than very short periods. The presence of2% w/v PEG 8000 in the buffer results in a very smooth-flowing resin suspension, and virtually eliminatesadhesion of resin particles to plastic surfaces. This is especially helpful when loading the column.113Bound proteins were eluted from the column with degassed 2-mL fractions of CEB-300, CEB-500,CEB-1000, and CEB-2000 (which contained 0.30, 0.50, 1.0, and 2.0 M KC1, respectively). Bound proteins wereeluted from the column using a flow rate of —0.10 mLlmin, and this flow rate was controlled by negative airpressure. The column was sealed with a slip-on, natural rubber septum (size 14), and filter-sterilized air wasintroduced above the liquid phase using a clamped syringe needle and line connected to a peristaltic pump.After capping of the Luer outlet, the line was broken and the needle was left in the septum whenever the septumwas fitted on or removed from the top of the column (i.e., to prevent disruption of the column bed when addingthe aliquots of CEB). Eluted fractions (2.0-mL) were collected directly into 2-mL ultrafiltration devices(Ultrafree-CL low protein-binding cellulose; 10,000 nominal Mr limit; Millipore UFC4LGC25), and thesefractions were stored on ice. After use the column was washed twice with two 12-mL aliquots of degassedCEB-2000 and then stored in the dark at 4 °C.Each eluted fraction was concentrated to a volume of —0.25 to 0.30 mL by centrifugation at 4 °C at2,400 g using an SS-34 rotor and RC-5 centrifuge. The concentrated fractions were transferred to 0.5-mi.microcentrifuge tubes, frozen quickly in a dry ice/ethanol bath, and then stored at -70 °C.D. IMMOBILIZED DNA AS A SUBSTRATE FOR ENZYME-CATALYZED REACTIONS.Standard DNA-immobilization reactions were performed using buffer TC and linearized pUC 19 DNA.Immediately after the initial stop reaction the remaining supernate (—1 mL) above aliquots of resin (40 iL,containing 1.0 mg/mL of immobilized DNA) was removed by aspiration and each aliquot was washed two timeswith 2.0 mL of SB. Each resin aliquot was then washed three times with 2.0 mL of TEP pH 7.4 (TE pH 7.4containing 2% w/v PEG 8000). After the final wash the resin was pelletecl, the supernate was removed byaspiration, 40 IJL of TEP pH 7.4 was added to each resin aliquot, and the aliquots were pooled. This “stocksuspension” was assumed to contain 50% v/v resin in TEP and hence —0.5 mg of DNA per mL of suspension.Restriction endonuclease reactions (50 j.tL final volume in microcentrifuge tubes) contained 10 .tg offree or immobilized DNA, 20 U of either ApaL I or Ssp I, and the appropriate lx reaction buffer. The ApaL Ireactions also contained 0.10 mg/niL of acetylated (nuclease-free) BSA. The restriction endonuclease reaction114buffers and BSA were supplied as lOx and lOOx stock solutions by the vendor, respectively. The stocksuspension of resin was resuspended gently but thoroughly prior to the removal of each aliquot. For thereactions which used immobilized DNA, all of the reaction components except the resin suspension were mixedgently, pelleted briefly, and then the resin suspension (20 jiL of 50% v/v in TEP, —0.5 mg of DNA per mL ofsuspension) was added by carefully expelling the suspension from the pipette tip directly into the liquidcollected at the bottom of the tube. After the resin suspension was added the reaction was mixed by gentlypipettmg the solution up and down a few times. After this step, do not pellet the resin prior to completion ofthe reaction. Reactions (3 h at 37 °C) were mixed by rotation at —2 rpm in a preheated incubator. However,mixing by rotation was later found to be unnecessary.When the restriction endonuclease reactions were completed, the tubes were very gently vortexed andthen centrifuged briefly. Equal aliquots (10 iL) of each digest were removed to microcentrifuge tubes, stopped,and then analyzed by electrophoresis on a 1.5% agarose gel in the presence of EtdBr using standard procedures.The unused portion of the TEP-washed resin should be either stored temporarily (if it is to be usedwithin —1-2 days), or it should be washed with SB or some other buffer suitable for longer term storage of theresin. The resin should be stored in the dark at 4°C.115CHAPTER 2.3. RESULTSA. THE IMMOBILIZATION OF DNA ON ACYL HYDRAZIDE-DERIVATIZED SOLiD SUPPORTS.1. Development of the microscale DNA-immobilization protocol.A prerequisite for the microscale DNA-immobilization reactions was the preparation of a large amountof high purity DNA to serve as a standard immobilization substrate. High purity DNA was required because theassay for immobilized DNA was based on the absorbance of 260-nm UV light. Contaminating proteins or RNAin the substrate DNA would compromise these assay results. Very large quantities of substrate DNA wererequired in order to examine a variety of reaction parameters in a statistically meaningful way. Variousparameters were examined in order to optimize the efficiency of the reaction, and thereby minimize exposure ofthe substrate DNA to the moderately acidic conditions that are essential for the reaction of cytosine derivativeswith acyl hydrazides. (Hayatsu & Ukita, 1964; Kikugawa et al., 1967a; Gal-Or et al., 1967).Watson-Crick hydrogen bonding strongly inhibits cytosine transamination reactions (Draper, 1984;Hayatsu, 1976a, 1976b; Hayatsu & Ukita, 1966; Shapiro et al., 1973). This led me to select Kpn IlinearizedtpUC 19 plasmid DNA as a standard immobilization substrate, as these DNA fragments have 3’-deoxycytidylateresidues at the end of their single-stranded caudemers. Each preparation of DNA was given a lot number (Ki,K2, etc.) and stored in aliquots so that results could be compared from each set of reactions that used the samelot of substrate DNA.In the DNA-immobilization protocol small portions of resin were washed with a coupling bufferprepared at 1 .25x its final concentration in the immobilization reaction (1.25x CB). Affi-Prep HZ, acommercially available, acyl hydrazide-denvatized, poly(methyl methacrylate)-based resin, was examinedinitially. The washed resin was used to make a 50% v/v suspension, and then microscale DNA-immobilizationreactions were carried out in quadruplicate using 80 j.LL of 50% v/v resin suspension and 20 iL of —2 mg/mLsubstrate DNA per reaction (Fig. 20). After incubation at 37 °C, each 0.100-mL reaction was stopped with 2.00mL of 0.5 M phosphate (sodium salt, pH 7.0), and then the uncoupled DNA was assayed spectrophotometrically.t The recognition sequence of the restriction endonuclease Kpn I is (G G TA C C ) (Roberts & Macelis, 1993).116The average apparent yield of immobilized DNA was calculated as mg of DNA per mL of resin.Although the immobilization reactions were performed in quadruplicate, the value of the standard deviation foreach set of reactions is not reported because these values would be misleading. The standard deviation for eachquadruplicate set of AA260 data was very low (<±0.005 or < 2%). The largest source of error in the averageapparent yield of immobilized DNA arose from the preparation of the 50% v/v resin suspension, not the zA260data. The “50% v/v” resin suspension could be dispensed with precision (based on the AA260 data), but theactual volume of resin in each 80 I.IL aliquot of the “50% v/v” resin suspension was less than the assumed valueof 40 iL (400 mg of resin and 400 iL of 1 .25x CB typically gave —760 jiL of degassed suspension). Hence theaverage apparent yield of immobilized DNA was systematically underestimated, probably by about 5%.2. Formulation of the coupling buffer.a. The hydrogen-ion buffer and its counterion.The reaction of cytosine derivatives with acyl hydrazides is sensitive to the hydrogen-ion concentration,and has a very sharp optimum at pH 4.2 (Hayatsu & Ukita, 1964; Kikugawa et al., 1967a; Gal-Or et al., 1967).Various hydrogen-ion buffers with one or more pKa values near 4.2 were therefore examined for their effect onthe immobilization reaction. These buffers were adjusted to pH 4.20 with KOH, with the exception of4-aminobutanoic acid (which was adjusted to pH 4.20 with HC1).Diethylenetriamine pentaacetic acid (DTPA) is a polyprotic weak acid with pKa values of 1.8, 2.6, 4.3,8.6, and 10.6 (Dean, 1992). If the DTPA in the coupling buffer was adjusted to pH 8.3, no DNA wasimmobilized on the resin (Table 5). However, if the DTPA in the coupling buffer was adjusted to pH 4.20,about 360 .tg of DNA was immobilized per mL of resin after 16 h at 37 °C. Formic, oxalic, and citric acids allgave very similar results to DTPA (Table 5). The results obtained with formic acid were the same whether itsfinal concentration in the coupling buffer was 10 or 30 mM, suggesting that within this range the concentrationof the weak acid had very little effect on the efficiency of the reaction.Tricarballylic acid (TCBA) is structurally identical with citric acid, except that it lacks the hydroxylgroup at C-3 (see Table 7). TCBA was almost twice as effective as citric acid in catalyzing the immobilization117reaction (Table 5). Another surprisingly effective buffer was 4-aminobutanoic acid (GAB A). In this case they-amino group of GABA may assist in the catalysis of the reaction. This may be analogous to the behaviour ofthe amino group during the catalysis by cysteainine of hydrogen-deuterium exchange at C-5 in uridine (Heller,1968; see also Hayatsu, 1976a).Table 5. Influence of wealdy acidic buffers on the covalent immobilization of DNA on Affi-Prep HZ.DNA immobilized (b)Coupling buffer (a) Comments (mg/mL resin)(CB)(1.1 = 100% yield)DTPA (diethylenetriamine-pentaacetic acid) 1 .25x CB adjusted to pH 8.3 0DTPA 0.36Formic acid 0.38Formic acid 30 mM final concentration 0.39Oxalic acid 0.35TCBA (tricarballylic acid) 0.76Citric acid 0.41GABA (4-ammobutanoic acid) 0.70Coupling buffers (1 .25x) were adjusted to pH 4.20 with KOH (HC1 for GABA), filter-sterilized, then used towash small portions of resin. Four aliquots of each washed resin suspension (80-pL aliquots, 50% v/v, lot36737) were distributed to cryotubes, DNA was added (20 .tL per tube, 2 mg/mL in 10 mM KC1, lot Ki), andreactions were mixed by rotation for 16 h at 37 °C. Stop buffer (2.00 mL of SB; 0.5 M phosphate, sodium salt,pH 7.0) was added and mixing continued 2 h as before. The resin was then pelleted and uncoupled DNA wasassayed spectrophotometrically.(a) Except as noted, final concentrations of weak acid and PEG 8000 in the coupling reactions were 10 mM and10% w/v, respectively. No attempts were made to maintain constant ionic strength.(b) Average (n=4) apparent yield for the immobilization of Kpn I-linearized, pUC 19 plasmid DNA (lot Ki).In the next series of DNA-immobilization reactions TCBA was used to maintain the pH in couplingbuffers that were adjusted to pH 4.20 with various bases (Table 6). Most of these bases supplied a counterionthat decreased the efficiency of the reaction compared with K+. However, the cyclic diamine piperazine118significantly increased the efficiency of the reaction, and it did so whether TCBA or citric acid was used as a pHbuffer (Table 6). The use of the piperazine analogue DABCO as TCBA’s counterion reduced the efficiency ofthe immobilization reaction by more than half compared with piperazine (Table 6). DABCO has two tertiaryamines instead of the two secondary amines in piperazme (see Table 7). At pH 4.20 the most predominant formof DABCO (pKas = 2.9, 8.6; Dean, 1992) in aqueous solution carries only a single positive charge, whereasunder the same conditions the most predominant form of piperazine (pKaS = 5.3, 9.8; Dean, 1992) carries twopositive charges. Compared with the DABCO monocation, the piperazinium dication in the pH 4.20 couplingbuffer would have been present at about half the concentration, and it would also have been more stronglyattracted to the DNA. Table 7 provides a summary of the structures, pKa values, and relative efficiencies of thevarious hydrogen-ion buffers and bases that were used to immobilize DNA on Affi-Prep HZ.Table 6. Influence of tricarbaflylic acid’s counterion on the immobilization of DNA on Affi-Prep HZ.DNA immobilizedBase added to adjust pH Comments (mg/mL resin)(1.1 = 100% yield)NaOH 0.51NH4O 0.41TRIS 0.46TRIS 1.25x CB adjusted to pH 8.3 0BIS-TRIS 0.51DABCO (1 ,4-diazabicyclo[2.2.2joctane) 0.42Piperazine 0.94Piperazine TCBA replaced by citric acid 0.70Reactions (n=4) were carried out as described in Table 5, except as noted. TCBA and PEG 8000 (10 mM and10% w/v final concentrations, respectively) were present in all reactions (except as noted). Coupling buffers(1.25x) were adjusted to pH 4.20 with the indicated bases. Kpn I-linearized, pUC 19 DNA (lot Ki) wasimmobilized on the resin (lot 36737) for 16 h at 37 °C.Table 7. Structure and relative efficiency of buffers used to immobilize DNA on Affi-Prep HZ.Hydrogen-ion buffer Base added to adjust pH Relative[pKa values] efficiencyCOOH COOHHOOCNCOOH KOH 0.38COOHDiethylenetriamine-pentaacetic acid (DTPA)[1.8, 2.6, 4.3, 8.6, 10.6]0‘I.H— CKOH 0.40Formic acid[3.8]HO 044 —C— C0” “OH KOH 0.37Oxalic acid[1.3, 4.3]HOOC(COOHCOOH KOH 0.80Tricarballylic acid (TCBA)[3.7,4.9,6.4]HOOC”fCOOH KOH 0.43COOH Piperazine 0.74Citric acid[3.1,4.8,6.4]H N’COOH2 HC1 added to adjust pH 0.754-aminobutanoic acid (GABA)[4.0 (COOH), 10.6]119120Hydrogen-ion buffer Base added to adjust pH Relative[pKa values] efficiencyTCBA NaOH 0.54[15.7]TCBA NH3 0.44[9.212HOTCBA 0.49OH OHTris(hydroxymethyl)aminomethane (TRIS)[8.1](OHTCBA HO’>(” OH 0.54N, ‘v ..Bis(hydroxyethyl)tris(hydroxymethyl)ammomethane (BIS-TRIS)[6.5]TCBA N I N 0.451,4-Diazabicyclo[2.2.2]octane (DARCO)[2.9, 8.6]/-TCBA HN NH 1.0\—/Piperazme[5.3, 9.8]The structures shown are the un-ionized forms of the various acids and bases used. All hydrogen-ion bufferswere present at a final concentration of 10 mM. The pKa values shown in square brackets (which are for theconjugate acids of the bases shown) are taken from a recent summary (Dean, 1992). Relative efficiency is forthe immobilization of Kpn I-linearized pUC 19 DNA (lot K1) on Affi-Prep HZ (lot 36737) as described inTables 4 and 5.121b. Other components of the coupling buffer.The efficiency of the DNA-immobilization reaction was increased by the addition of lowconcentrations (30-50 mM) of supplementary KC1 to the coupling buffer (Table 8). However, if thesupplementary KC1 concentration in the coupling buffer was too high, the efficiency of the reaction decreasedsignificantly, and at or above 200 mM KC1, no DNA was immobilized at all. I suspect that at low concentrationsthe supplementary KC1 may have increased the efficiency of the reaction by significantly increasing theefficiency of mixing, as 30-50mM KC1 appeared to increase the fluidity of the resin suspension.Table 8. Influence of supplementary KC1 onthe immobilization of DNA on Affi-Prep HZ.DNA immobilizedFinal concentration (mg/mi. resin)of KC1 (mM) (a) (1.1 = 100% yield)0 0.9430 1.150 1.1100 0.76200 0400 0Reactions (n=4) were carried out as describedin Table 5, except as noted. TCBA and PEG8000 (10 mM and 10% w/v finalconcentrations, respectively) were present in allreactions. Coupling buffers (1 .25x) wereadjusted to pH 4.20 with piperazine. Kpn Ilinearized, pUC 19 DNA (lot Ki) wasimmobilized on the resin (lot 36737) for 16 h at37°C.(a) The DNA solution contributed a further2 mM KC1 in all cases.Table 9. Influence of PEG 8000 on theimmobilization of DNA on Affi-Prep HZ.DNA immobilizedFinal concentration (mglmL resin)of PEG 8000 (% w/v)(1.1 = 100% yield)10 0.6112 0.6614 0.6816 0.4118 0.1720 0.12Reactions (n=4) were carried out as describedin Table 5, except as noted. TCBA and KC1(10 and 50 mM final concentrations,respectively) were present in all reactions. TheDNA solution contributed a further 2 mM KC1.Coupling buffers (1.25x) were adjusted to pH4.20 with piperazine. Kpn I-linearized, pUC 19DNA (lot Ki) was immobilized on the resin(lot 36737) for 4 h at 37 °C.122The next series of immobilization reactions examined the effect of polyethylene glycol (PEG) 8000 inthe coupling buffer (Table 9). This polymer served several roles. Addition of PEG to the coupling bufferresulted in a smooth-flowing resin suspension, and virtually eliminated adhesion of resin particles to plasticsurfaces. In addition, PEG is known to act as a volume exclusion agent (Lerman, 1971, 1973; Zimmerman &Minton, 1993). Each macromolecule of DNA is effectively excluded from the volume element occupied byeach macromolecule of PEG, which effectively increases the concentration of DNA in the solution. Underappropriate conditions the phenomenon of “macromolecular crowding” has been shown to increase the rate ofDNA renaturation, and the rates with which DNA interacts with other molecules (e.g., enzymes) (Amasino,1986; Sikorav & Church, 1991; Zimmerman & Harrison, 1987). However, care must be taken when using PEG,as under certain conditions it can precipitate DNA (Lerman, 1973; Lis & Schleif, 1975).Polyethylene glycol (with an average Mr of 8,000) at a final concentration of 12 to 14% w/v in thereaction gave the best results, but at higher PEG concentrations the efficiency of DNA immobilization decreased(Table 9). At this stage in the study the optimal reaction conditions included the following solutes at theindicated final concentrations: 10 mM TCBA, 52 mM KC1, and 14% w/v PEG 8000, with the 1 .25x couplingbuffer adjusted to pH 4.20 with piperazine. Using these conditions approximately 680 jig of Kpn I-linearized,pUC 19 DNA (60% of the total DNA used in the reaction) could be immobilized per mL of Affi-Prep HZ after4hat37 °C (Table 9).The next series of immobilization reactions examined the effects of the hydrogen-ion concentration,and the type and concentration of the cyclic polyamine used in the coupling buffer (Table 10). These reactionsused pUC 19 DNA that had been linearized with either Kpn I or Xba I restriction endonucleasest, and whichtherefore possessed either 3’- or 5’-deoxycytidylate residues, respectively, at the end of their single-strandedcauclemers. Both types of DNA gave very similar results (Table 10). As expected, if the pH was increasedabove 4.2 the efficiency of the immobilization reaction decreased significantly. No attempt was made toexamine the efficiency of the reaction at pH values lower that 4.2, as these conditions were considered to be toopotentially harmful to the DNA.t The recognition sequence of the restriction endonuclease Xba I is (T .1- C TAG A) (Roberts & Macelis, 1993).Table 10. Influence of critical cations on the immobilization of DNA on Affi-Prep HZ and Affi-Gel HZ.DNA immobilized (mg/mL resin)Coupling buffer (a) Comments (1.2 = 100% yield)(CB)Kpn I-linearized DNA Xba I-linearized DNATP l.25x CB adjusted to pH 4.40 0.74 0.67TP 0.93 0.85TC 1.0 1.0TC Affi-Gel HZ resin (30 °C) 1.1 1.1rr 0.18 0.23TC 0.12 0.10Reactions (n=4) were carried out as described in Table 5 with some minor modifications. Unless specifiedotherwise, Kpn I-linearized (lot K2; 2.42 mg/rnL) or Xba I-linearized (lot Xl; 2.38 mg/mL) pUC 19 DNA(20 j.il. of DNA in 10 mM KC1 added per reaction) was immobilized for 6 h at 37 °C on Affi-Prep HZ resin (lot417 lOB) or at 30 °C on Affi-Gel HZ resin (lot 43482B).(a) The final concentrations of the components in the coupling buffers examined are given below. The finalKC1 concentrations specified include 2 mM KC1 contributed by the DNA solution.“T” buffers: 8 mM TCBA 50 mM KC1; 12% w/v PEG 8000 (1 0.06 to 0.07 M)“TT” buffers: 16 mM TCBA 42 mM KC1; 12% w/v PEG 8000 (I— 0.06 to 0.07 M)The pH of the 1.25x CBs was adjusted to 4.20 (except as noted) with either piperazine (P) or cyclam (C).There was a slight increase in the efficiency of the immobilization reaction if piperazine was replacedby the cyclic tetraamine 1,4,8,1 1-tetraazacyclotetradecane (cyclam). The final, optimized formulation of thecoupling buffer was designated TC, and it contained the following solutes at the indicated final concentrations:8 mM TCBA, 50 mM KCI, and 12% w/v PEG 8000, with the 1 .25x coupling buffer adjusted to pH 4.20 withcyclam. Using buffer TC, 1.0 mg of Kpn I-linearized, pUC 19 DNA (80% of the total DNA used in the reaction)could be immobilized per mL of Affi-Prep HZ after 6 h at 37 °C (Table 10).The yield of immobilized DNA was slightly higher when Affi-Gel HZ, an agarose-based resin, wasused instead of Affi-Prep HZ. The reactions performed with Affi-Gel HZ were carried out at 30 °C because thistemperature is the upper limit recommended by the vendor for the use of this matrix. This resin was used to123124visualize the immobilized DNA by microscopy after staining of the gel beads with the DNA-specificfluorochrome Hoescht 33258 (see Fig. 21). All beads containing immobilized DNA showed strong fluorescentstaining (Fig. 21a). However, when a similar staining protocol was used with a 1:1 mixture of beads containingimmobilized DNA and beads that had been subjected to a mock immobilization reaction (without DNA),fluorescent staining was only observed with some beads but not with other beads (i.e., when the same field ofobservation was photographed using DIC and epi-fluorescence methods) (Fig. 21b).Table 11. Structure, relative efficiency, and inhibitory potential of piperazine and cyclam.Structure pKa values Relative efficiency Inhibitory potentialr-\HN NH 5.3, 9.8 1.0 0.81\-,1=0Piperazine/C NH HN 1.6, 2.4, 10.6, 11.6 1.1 0.89NH HN I=O.5M (KNO3)\—JCyclam(1,4,8,1 1-tetraazacyclotetradecane)The structures shown are the un-ionized forms of the bases, and the pKa values are for their conjugate acids(Dean, 1992; Micheloni et al., 1978). Relative efficiency is for the immobilization of Kpn I-linearized pUC 19DNA (lot K2) on Affi-Prep HZ (lot 41710B) at 37 °C using either buffer TP or TC as described in Table 10.The inhibitory potential was based on results obtained at 37 °C using the same DNA and resin, and was definedas the fractional reduction in the average (n=4) apparent yield of immobilized DNA when the finalconcentration of TCBA in the coupling buffer was doubled from 8 to 16 mM (using buffers TP and HP or TCand TTC as described in Table 10).125Figure 21. Microscopy of Affi-Gel HZ beads containing immobilized DNA.Two samples of Affi-Gel HZ beads (a and b; described below) were each washed three times with 2 mL ofhigh-salt assay buffer (2.0 M NaC1; 50 mM phosphate, sodium salt; pH 7.4; containing 100 ng/mL of theDNA-specific fluorochrome Hoescht 33258), and then examined by DIC (left panels) or epi-fluorescence (rightpanels) methods using a Zeiss 1M-35 microscope.(a) panels (L&R): Affi-Gel HZ beads (40 jiL) from a standard immobilization reaction (using pUC 19 DNA)were stopped with SB, washed twice with 2.0 mL of SB, and then treated further as described above. Both thefield of observation and its orientation are the same in the left and right panels.(b) panels (L&R): Affi-Gel HZ beads (40 p.L of each type) from a mock immobilization reaction (using KC1instead of DNA) and a standard immobilization reaction (using pUC 19 DNA) were stopped with SB, and thenmixed 1:1. The 1:1 mixture of beads (80 iL) was washed twice with 2.0 mL of SB, and then treated further asdescribed above. Both the field of observation and its orientation are the same in the left and right panels.(a)(b)126Hydronium ion is not the only positively charged ion whose concentration had a dramatic influence onthe efficiency of the immobilization reaction. Previously, when the (final) concentration of formic acid in thecoupling buffer was tripled from 10 to 30 mM, there was no effect on the efficiency of the immobilizationreaction (Table 5). However, if the (fmal) concentration of TCBA in buffer TC was doubled from 8 to 16 mM(to give buffer TTC) the efficiency of the immobilization reaction dropped precipitously (Table 10). BufferTC had twice the concentration of TCBA as buffer TC, but it also had twice the concentration of cyclam. Inthe formulation of buffer TrC, the KC1 concentration was lowered relative to its concentration in buffer TC, i.e.,from 50 to 42 mM, in an attempt to a minimize the difference in the ionic strengths of the two buffers.The difference between the results obtained using buffers TP and Ti? (where the cyclic polyamine inthe coupling buffer was piperazine) was very similar to the difference in the results obtained using buffers TCand TTC (Table 10). However, a closer look at the data indicated that just as cyclam was more efficient as acounterion for TCBA than piperazine (cf. the results with buffers TC and TP), cyclam was also more efficientthan piperazine at inhibiting the immobilization reaction when the TCBA concentration in the coupling bufferwas doubled from 8 to 16mM (Jable 11).3. The nature of the deoxycytidylate-acyl hydrazide linkage.Up to this point in the study I assumed that the major—if not the only—point of attachment of the DNAto the solid support was the terminal deoxycytidylate residue on either of the single-stranded caudemers of eachDNA fragment. This assumption was incorrect, at least for the immobilization of DNA using buffer TC (Table12; see also Fig. 23). Little if any effect on the relative efficiency of the immobilization reaction was observedwhen buffer TC was used to immobilize pUC 19 DNA that had been linearized with a restriction endonucleasethat generated either no caudemers (Ssp I or Sma I) or caudemers that lack deoxycytidylate residues (EcoR I).Previous reports had indicated that deoxycytidylate residues in double-stranded DNA do not react withacyl hydrazides, e.g., semicarbazide (Hayatsu, 1976b; Hayatsu & Ukita, 1966), nor are they susceptible tobisulfite-catalyzed transamination reactions (Draper, 1984; Hayatsu, 1976a; Shapiro et at., 1973). In theinterpretation of these studies, however, it should be noted that they all used high ionic-strength conditions, i.e.,high concentrations of bisulfite and transaminating reagent (acyl hydrazide or primary amine) to increase the127reaction rate. In this study the concentration of the acyl hydrazide in the reaction was fixed (and contributedlittle to the ionic strength of the solution), no bisuffite catalyst was required, and the use of higher ionic-strengthconditions was clearly inhibitory (Tables 7 and 9). Under low ionic-strength conditions, localized, transientopening of the DNA helix may have exposed single-stranded deoxycytidylates at points along the length of theDNA fragment, and this process may have been essential for the reaction to proceed to any significant extent.Transient, localized opening of the DNA double-helix was probably also favoured by the moderately acidicreaction conditions (Banville et at., 1986; Courtois et at., 1968; Gehring et at., 1993; O’Connor et at., 1981;Robinson et at., 1992; Smol’janinova et at., 1982; Zimmer & Venner, 1966; Zimmer et al., 1968).Table 12. Influence of caudemer sequence on the immobilization of DNA on Affi-Prep HZ.Restriction endonuclease Fragment termini Relative efficiency (a)XbaI pCpTpApGpA- 1.0TAvaI pCpCpGpGpG- 1.0CEcoRI pApApTpTpC- 1.0GAccI pCpGpApC- 0.90TpGSspI pApTpTp- 0.92TpApAp -SinaI pGpGpGp- 0.89CpCpCpPlasmid DNA (20 jiL of 1.8 mg/mL of pUC 19 DNA in 10 mM KC1 added per reaction) that had beenlinearized with the indicated restriction endonuclease was immobilized (n=4) for 6 h at 37 °C on Affi-Prep HZresin using buffer TC as described in Table 10. Restriction endonuclease specificities and linearized pUC 19fragment tennini are described in Roberts & Macelis (1993) and Yanisch-Perron et at. (1985).(a) Coupling efficiency is a ratio equal to the average apparent yield of immobilized DNA divided by theaverage amount of input DNA in each set (n=4) of immobilization reactions. The relative efficiency is thecoupling efficiency for each set of reactions divided by the coupling efficiency for the set of reactions thatused Xba I-linearized pUC 19 DNA.128--L]LJ +12345— —TRIS-HCI (100 mM, pH 8.0)Bisulfite (200 mM)Bisulfite (20 mM)Resin12345Figure 22. Bisulfite-catalyzed immobilization of DNA on polyacrythydrazido agarose.Polyacrythydrazido agarose was hydrated and then washed extensively with 30 mM citrate, sodium salt, pH 4.5(“citrate buffer”). Where indicated, an aliquot (1.0 mL of settled resin) of the washed resin or 1.00 mL of citratebuffer was added to a cryotube. Bisulfite (12 or 120 .L of a 2.0 M stock of freshly prepared sodium bisulfite,pH 4.2-4.3), or TRIS-HC1 (120 IlL of a 1.0 M stock, pH 8.0) was added as indicated; additional citrate bufferwas also added to adjust the volume to 1.12 mL in all cases (if necessary). An aliquot (80 jiL) from a stocksolution of digested, purified, double-stranded plasmid DNA (consisting of two Apa I fragments of —2,700 and—350 bp) was added to each tube. The 1.2-mL reactions were mixed by rotation at 50 rpm in a pre-warmedincubator for 18 hours at 37 °C, and then 0.30 mL of 5.0 M NaCl was added and the reactions were mixed afurther 5 mm. The resin was pelleted, 0.75 mL of the supernate from each reaction was removed tomicrocentrifuge tubes, and then the DNA was precipitated overnight at 4 °C with 1 vol of 2-propanol. TheDNA was pelleted, all traces of the supernate were removed (no rinse), and then the DNA pellets were air-driedbriefly. The DNA was resuspended in 36 pL of TE pH 8.0, and then 12 .tL aliquots from each reaction wereanalyzed by agarose gel electrophoresis (0.6% gel) in the presence of EtdBr using standard methods.Bisuffite or TRIS-HC1 were present at the indicated fmal concentrations.129Although no bisulfite catalyst was required for the immobilization of double-stranded DNA onAffi-Gel HZ or Affi-Prep HZ, this catalyst may be used to increase the rate of immobilization. At the beginningof this study I used a different resin (polyacrythydrazido agarose from the Sigma Chemical Co.), and underconditions where essentially no DNA was immobilized on this resin (using 30 mM citrate, sodium salt, pH 4.5,as a coupling buffer) (Fig. 22, cf. lanes 1 and 2), the addition of as little as 20mM bisulfite to the immobilizationreaction led to a dramatic increase in the amount of DNA immobilized (Fig. 22, lane 3). These data also indicatethat the mechanistic basis of the immobilization reaction is probably transamination at transientlysingle-stranded deoxycytidylates. These data are inconsistent with the hypothesis that the immobilizationreaction involves the formation of acyl hydrazones at apurinic sites in the DNA (see also the Discussion Sectionfor additional consideration of this hypothesis).The plasmid pUC 19 is 2,686 bp long (Yanisch-Perron et al., 1985). Although linear DNA of thislength could be immobilized with satisfactory efficiency on acyl hydrazide-derivatized solid supports, under thesame reaction conditions duplex oligonucleotides reacted very poorly (Table 13). Only 140-250 jig of a duplex51-bp HSE probe (14 to 24% of the DNA used in the reactions) could be immobilized per mL of resin usingbuffer TC. Unfortunately, I did not have enough of the three types of probe (hse-A•hse-T, hse-R•hse-Y, andnonphosphorylated hse-R•hse-Y) to examine the immobilization efficiencies of each type of DNA on bothAffi-Prep HZ and Affi-Gel HZ. The greater immobilization efficiency of the 51-bp HSE probe hse-A•hse-T onAffi-Gel HZ (cf. the results with hse-R•hse-Y on Affi-Prep HZ) probably reflects the superior reactivity orcapacity of this matrix. For the reasons discussed above, the caudemers of the duplex oligonucleotide probes(Fig. 3) probably had little if any effect on the yield of immobilized DNA.For very small DNA fragments the efficiency of the DNA-immobilization reaction appeared to increasewith an increase in the length of the DNA. If hse-A•hse-T was first polymerized using T4 DNA ligase and thenimmobilized to the resin (using the same conditions employed with the 51-bp monomer), the efficiency of thereaction doubled (Table 13). It should be noted that the large-scale ligation reaction used to polymerizehse-A•hse-T was inefficient, and yielded a mixture of fragments with an average length of only -l50 bp. Hencein the preparation of sequence-specific DNA-affinity chromatography resins (e.g., see Kadonaga, 1991;130Kadonaga & Tjian, 1986) the efficiency of the ligation reaction (i.e., the average fragment length) would in turnaffect the efficiency with which the polymerized DNA could be immobilized on an acyl hydrazide-derivatizedsupport.Table 13. Preparation of DNA-affinity chromatography resins.Resin DNA (a) Length (bp) DNA immobilized Efflciencyhse-R•hse-Y 51 0.14 14hse-R•hse-Y 51 (nonphosphorylated) 0.18 17Affi-Prep HZpoly(dI-dC).poly(dI-dC) —8,600 1.7 85poly(dI-dC)’poly(dI-dC) —1,200 1.6 81hse-Ahse-T 51 0.25 24poly(hse-A.hse-) -150 0.52 49Affi-Gel HZpoly(dI-dC).poly(dI-dC) -.8,600 1.7 89poly(dI-dC)•poly(dI-dC) —1,200 1.8 92Reactions (n=4) were performed as described in Table 10 using buffer TC. DNA samples were immobilized for6 h at 37 °C on Affi-Prep HZ resin (lot 41710A) or at 30 °C on Affi-Gel HZ resin (lot 43482B).(a) Specifications of the DNA samples are given below. Oligonucleotides were chemically phosphorylated atthe 5’ end except as noted.duplex HSE probes: —29 A260 units/mg —2.0 mg/mL;poly(hse-A•hse-T): -.29 A260 units/mg —2.0 mg/mL;lot I-i poly(dI-dC)•poly(dI-dC): 10.0 A260 units/mg —3.6 mg/mL;lot 1-2 poly(dI-dC)•poly(dI-dC): 12.7 A260 units/mg —3.6 mg/mL.The synthetic, alternating copolymer poly(dI-dC)•poly(dI-dC) could be immobilized on acyl hydrazidederivatized resins very efficiently (Table 13). Two different lots of this DNA, with average fragment sizes of1,200 or 8,600 bp (as specified by the vendor), were examined. The results indicated that within this range theDNA fragment size had no effect on the yield of immobilized poly(dI-dC)•poly(dI-dC). These results alsosuggested that the observed yields of immobilized pUC 19 DNA on acyl hydrazide-derivatized resins (1.0-1.1mg/mL) may have been even greater if higher concentrations of DNA were used in the immobilization reactions.1314. The stability of the deoxycytidylate-acyl hydrazide linkage.The assay for the stability of the linkage was very simple. Various samples of resin (containingquantitated amounts of immobilized DNA) were washed extensively and then resuspended with a knownvolume of stop buffer (SB). Sodium fluoride was then added to a fmal concentration of 50 mM to inhibitmicrobial growth. After 10 months of storage at 4 °C any DNA that had been released into the supernate (i.e.,lost from the resin) was assayed specirophotometricafly.Table 14. Loss of immobilized DNA after ten months.DNA initially immobilizedResin DNA lost afterten monthsType Length (bp) mg/mL resin (%)Kpn I-linearized pUC 19 (lot K2) 2,686 1.0 41Affi-Prep HZ Xba I-linearized pUC 19 (lot Xl) 2,686 1.0 40Poly(dI-dC).poly(dI-dC) (lot I-i) —8,600 1.5 62Kpn I-linearized pUC 19 (lot K2) 2,686 1.1 31Affi-Gel HZ Xba I-linearized pUC 19 (lot Xl) 2,686 1.1 31Poly(dl-dC).poly(dI-dC) (lot 1-1) —8,600 1.8 90Aliquots (n=4) of resin (40 .tL, containing quantitated amounts of DNA that had been immobilized as describedin Table 10) were washed extensively with SB and then stored under 2.16 mL of SB (containing 50 mM NaF) at4 °C for 10 months. After the period of storage was over, the resin aliquots were equilibrated to 20-22 °C,mixed by rotation, pelleted, and then the supernate was assayed spectrophotometrically. The resins examinedwere Affi-Prep HZ (lot 41710B) and Affi-Gel HZ (lot 43482B).The results obtained with Kpn I- and Xba I-linearized pUC DNA were identical. After 10 monthsstorage at 4 °C theit were losses of 40% and 30% of the initially immobilized pUC 19 DNA from Affi-Prep HZand Affi-Gel HZ, respectively (Table 14). The losses of poly(dI-dC)•poly(dI-dC) from these resins were muchhigher. After 10 months storage at 4 °C more poly(dI-dC).poly(dI-dC) was lost from Affi-Gel HZ (90%) thanfrom Affi-Prep HZ (60%), the opposite pattern to that observed with linearized pUC 19 DNA. Unfortunately,lot 1-2 poly(dI-dC)•poly(dI-dC) (with an average fragment length of only 1,200 bp) was not available at the time132this experiment began. Hence it is not clear if any relationship exists between the size of the immobilized DNAfragment and the stability of the deoxycytidylate-acyl hydrazide linkage. Poly(dI-dC)•poly(dI-dC) may beunusually reactive, and the linkage between this type of DNA and the resin may be unusually unstable, becauseof the propensity of alternating copolymers containing deoxycytidylates to adopt unusual structures (via theformation of cytosine’cytosine cation base pairs) under the moderately acidic conditions used during theimmobilization reaction (e.g., see Gray et al., 1980; Robinson et a!., 1992). Alternatively, the much greaterlosses of poly(dI-dC)•poly(dI-dC) from the resin may have been due to the shedding of hairpin (unimolecular)duplexes formed by the complementary, unimmobilized strands of (bimolecular) duplex molecules that had beenimmobilized on the resin through a single covalent bondt. The latter explanation is consistent with theobservation that a greater percentage of poly(dI-dC)•poly(dI-dC) was lost from Mn-Gel HZ than Mfi-Prep HZ.The agarose-based resin is the more porous of the two resins examined, and hence poly(dI-dC).poly(dI-dC)immobilized within the pores of this resin would have relatively fewer steric constraints to the formation andloss of unimolecular hairpin duplexes.B. IMMOBILIZED DNA AS AN AFFINITY CHROMATOGRAPHY LIGAND.Poly(hse-A.hse-T) DNA was immobilized on 0.80 mL of Affi-Gel HZ to serve as an HSE-specificDNA-affinity chromatography matrix. An A-protocol nuclear extract was prepared from heat-shocked eggs andused to determine if the putative ceHSBFm activity in this extract might be retained by the poly(hse-A.hse-T)minicolumn. No attempt was made to fractionate the A-protocol nuclear extract prior to chromatography on theaffinity column because the amount of starting material from which the extract was prepared was very limited(4 g of eggs). My primary objective was to determine if poly(hse-A.hse-T) DNA immobilized on Affi-Gel HZwas able to function as a sterically accessible affmity chromatography ligand.The A-protocol nuclear extract was adjusted to —150 mM KC1, insoluble proteins were removed, andthen the solution was applied to the 0.8-mL HSE minicolumn. The column was washed with CWB and thenbound proteins were eluted with 2.0-mL aliquots of CEB containing 0.3, 0.5, 1.0, or 2.0 M KC1. These fractionst I gratefully acknowledge that this explanation was suggested by Prof. C. R. Astell.133were concentrated and then used in EMSAs to determine the level of the putative ceHSBFm activity in eachfraction. SDS-PAGE was also performed to examine the protein content of each fraction.By comparing the relative intensities of the Rf 0.4 and Rf 1.0 signals from lane 1 of Fig. 14b and lane 3of Fig. 23a, I estimated that only25%t of the putative ceHSBFm activity in the nuclear extract was retained bythe poly(hse-Ahse-T) minicolumn and recovered in the 1.0 M KC1 fraction. The recovery of the putativeceHSBFm activity would probably have been better if the bulk of the nonspecific DNA-binding activities wereremoved by a prior fractionation step. Another factor which probably impaired the performance of the columnwas the small average DNA fragment size (—150 bp) of the poly(hse-A•hse-T) ligand.Figure 23. EMSA and SDS-PAGE analyses of HSE-DNA-affinity column fractions.Heat-shocked nematode eggs (4 g) were used to prepare 2.0 mL of nuclear extract using the A-protocol. Thisextract was adjusted to —150 mM KC1, 0.2% w/v 0G. and 0.2 mg/mL pdN in a final vol of 5.0 mL (with theextract supplying adequate pH-buffering and —140 mM of KC1). Insoluble proteins were removed and then thesolution was applied to an HSE-DNA-affinity minicolumn [0.80-mL bed volume; 0.42 mg of poly(hse-A.hse-T)immobilized per mL of Affi-Gel HZ resin; prupared as described in this thesis]. Chromatography was performedat 20-22 °C. The column was washed with 10 mL of CWB plus 0.2 mg/mL pdN and then 10 mL of CWB.Bound proteins were eluted with 2.0-mL aliquots of CEB containing 0.3,0.5, 1.0, or 2.0 M KC1, and the elutedfractions were kept on ice. Each eluted fraction was concentrated at 4 °C to about 0.25-0.3 mL, quick-frozen,stored at -70 °C, and then aliquots of these concentrated fractions were analyzed later as indicated below.(a) EMSA analysis. Aliquots (2 jiL or —0.8%) of each fraction were used in DNA-binding reactions (30 mm at20-22 °C, 20 .tl final vol, 0.5x BHE as the binding-reaction buffer, 6% w/v sucrose, 0.2 ng=6 fmol of a[32P1-labeled HSE probe per reaction). The estimated concentration of KC1 in each binding reaction wasadjusted to 150-200 mM (using CEB-500) as required. PAGE (using KGBM) separated the free probe(Rf=1.0, indicated on the left) from the less-mobile complex (Rf= 0.4). Only 9 j.LL of each reaction wasloaded (or —0.4% of the activity of each fraction). The gel was 4.5%T, 2.O%CBis and was prerun 0.5 h andrun 6 h at 40 V. The dried gel was autoradiographed for 14 h at -70 °C using one intensifying screen.(b) SDS-PAGE analysis. The high-salt buffer in aliquots (20 iL or —8%) of each fraction was exchanged forSTE at 4°C and then each fraction was concentrated to —20 iL using a microcenthfuge-driven ultrafiltrationunit (10,000 nominal Mr limit). Samples were analyzed by discontinuous SDS-PAGE (Laemmli, 1970)using a commercial apparatus (Hoefer, model SE 600) and standard conditions. The gel was 1.5-nun thick(cf. c below) and the separating gel was 10%T,3.%CBis. A commercial set of silver-stain high-Mrstandards (m) and a “Silver-Stain Plus” kit (Bio-Rad) were used according to the vendor’s instructions.(c) SDS-PAGE analysis. The high-salt buffer in the remainder (230-280 liL or —90%) of each fraction wasexchanged for STE at 4 °C and then each fraction was concentrated to —20 .LL as described in (b). Sampleswere then analyzed as in (b), except the gel was 0.75-mm thick.t I arbitrarily defined units (U) of retention in terms of a standard 10-p.L DNA-binding reaction (as describedin Fig 14b, lane 1) containing 3 fmol of labeled HSE probe, and set 1 U = 10% retention in the Rf 0.4 complexof the combined Rf0.4 and Rf 1.0 radioactivity. From lane 1 of Fig. 14b I estimated that 4 iL of extract gave60% retention (6 U), hence 2.0 mL of extract should have yielded —3000 U of retention. From Fig. 23a Iestimated that 1 jiL of the 1.0 M KC1 fraction would have given 30% retention (3 U) in a standard 10-.iLDNA-binding reaction, and hence this fraction (250-300 .iL) contained -750-900 U of retention.Columnfractions0.30.51.02.0I:lii.(m=sizemarkers)EMSAColumnfractionsIbiSDS.PAGEidlal00.41.0SDS-PAGEColumnfractions0.3I0.51.02.0m1ooo200116976645M10002001169766451234mm‘1234mm135It had previously been observed that in some cases the Rf 0.4 signal can be resolved into a closelyspaced doublet (see Fig. 1 1). A comparison of lanes 2-4 in Fig. 23a indicates that the Rf 0.4 signal in lane 3 mayalso be a doublet, and that the same signal in lanes 2 and 4 correspond to the upper and lower bands of thisdoublet, respectively. Thus it appeared that the protein that gave rise to the upper band of the doublet eluted atlower salt concentrations (0.5-1.0 M KC1), whereas the protein that gave rise to the lower band of the doubleteluted at higher salt concentrations (1.0-2.0 M KC1). It is unclear if more than one protein or related fonns ofthe same protein (i.e., a discrete proteolytic degradation product) gave rise to the two bands of the doublet.Interpretation of these results is difficult, however, because in most of the EMSAs performed using theunfractionated A-protocol nuclear extract the Rf 0.4 signal appeared as singlet. Furthermore, the columnfractions used for the EMSAs shown in lanes 2-4 in Fig. 23a contained very different concentrations ofcontaminating non-specific DNA-binding proteins, and the binding reactions themselves were performed usingslightly different concentrations of KCI (but in all cases in the range of 100-200 mM).The SDS-PAGE analysis of the protein content of each fraction showed that most of the proteins thatbound to the immobilized poly(hse-A.hse-T) DNA were eluted with 0.5 M KC1 (Fig. 23a, b). It was impossibleto detect any protein bands in the 1.0 and 2.0 M KC1 fractions in the initial analysis, so the remaining contents ofeach fraction were concentrated further and then analyzed on a thinner (0.75- vs. 1.5-mm) SDS-PAGE gel.Unfortunately, it was difficult to quantitatively recover each concentrated fraction from the disposablemicroconcentrator (due to the small size of each of the final volumes). The percentage of each fraction(nominally 90%) that was actually analyzed was probably 75 ±15%, with the percentage of the 1.0 M fractionanalyzed being at the low end of this range. It was therefore difficult to compare the intensities of thesilver-stained protein bands in Fig. 23c.In sequence-specific DNA-affinity chromatography, the eluted fraction that is found to contain thesequence-specific DNA-binding activity of interest is frequently adjusted to the appropriate salt concentrationand then rechromatographed on the affmity column (Kadonaga, 1991; Kadonaga & Tjian, 1986). The observedcomplexity of the protein profile of the 1.0 M KC1 fraction (i.e., after only one affinity chromatography step)was therefore not surprising. The spectrum of low-Mr proteins in this profile is similar to that observed in the1361.0 M KC1 eluate after the initial affinity chromatography of S. cerevisiae HSF (see Fig. 5 in Sorger & Petham,1987).Notwithstanding the caveats mentioned above, any candidate band in lane 3 of Fig. 23c representing theputative ceHSBFm protein would have to be absent from the 0.3 M KC1 fraction, and be more prominent in the1.0 M KC1 fraction than the 0.5 or 2.0 M KC1 fractions. The only band in the 1.0 M KC1 fraction that appearedto satisfy these criteria is the band that migrated just above the 66,000 Mr marker (Fig. 23c). This particularband is noteworthy because the Mr values of several HSF isolates are between 65,000-70,000 (Carmo-Avideset al., 1990; Kroeger et al., 1993; Nakai & Morimoto, 1993). Perhaps more importantly, higher Mr HSF isolates(from D. melanogaster and S. cerevisiae; Sorger & Petham, 1987; Wiederrecht et at., 1987; Wu et al., 1987)have been shown to give rise to a discrete proteolytic degradation product with an Mr in this range.C. IMMOBILIZED DNA AS A SUBSTRATE FOR ENZYME-CATALYZED REACTIONS.Sequence-specific DNA-affinity chromatography is a very important technique, and Affi-Gel HZappears to have potential to serve as a convenient and safe alternative to CNBr-activated Sepharose for thepreparation of DNA-affinity columns. However, covalently immobilized, unmodified, double-stranded DNAalso has the potential to serve as a substrate for a variety of physical, chemical or enzymatic reactions. The useof such DNA in enzyme-catalyzed reactions would be particularly useful, as applied enzyrnology is thefoundation of recombinant DNA technology. Unfortunately, solid-phase applications of these techniques wouldin most cases require that both the linkage and the parent support be able to tolerate elevated temperatures.Agarose-based resins would be unsuitable for such applications, as their recommended maximum operatingtemperature is 30 °C (as stated in Bio-Rad’s product information data for Affi-Gel HZ).• Linearize pUC 19 DNA with either Kpn I or Xba I.• Immobilize each type of DNA on Affi-Prep HZ.• Set up restriction digests with equal amounts of DNA.• Cut free (F) or immobilized (I) DNA with ApaL I or Ssp I.• Analyze digest aliquots by agarose gel electrophoresis.137Xba ITLCTAGAKpnlGGTACL-CApaL IGTGCACSspIAATATTmlApaL I SspI2345678FragmentApaL IKpn XbaI 1246I 702 697I 497241 2461234size (bp)SspIKpn Xba2083 120781603 6085678Figure 24. Restriction endonuclease digestion products obtained using immobilized DNA.The published DNA sequence of pUC 19 (Yanisch-Perron et al., 1985) was used to calculate the sizes of theexpected DNA fragments from the digests (right) and the DNA markers (m; 2,686; 1,444; 736; and 476 bp) forthe agarose gel (left). Restriction endonuclease recognition sequences are from Roberts & Macelis (1993).Kpn XbamFI FlKpn XbaFIFI138Affi-Prep HZ is a poly(methyl methacrylate)-based chromatography resin that possesses highmechanical strength as well as resistance to pH and temperature extremes, properties that are very desirable foraji HPLC support. Kpn I- or Xba I-linearized pUC 19 DNA was immobilized on Affi-Prep HZ and then used asa substrate for one of two restriction endonucleases, ApaL I or Ssp I. When these and the same reactionsperformed in solution were analyzed by agarose gel electrophoresis, the number and sizes of the digestionproducts were the same for comparable reactions (Fig. 24). However, the relative EtdBr-staining intensities ofthe digestion products differed when comparing the same reactions performed in solution or using immobilizedDNA (cf. the “I” and “F” lanes in Fig. 24).These restriction enzyme digest results not only indicated that DNA immobilized on Affi-Prep HZ maybe a suitable substrate for a variety of enzymatic reactions at elevated temperatures. These results (which werereproducibly obtained on two separate trials) also provided some information about the nature of thedeoxycytidylate-acyl hydrazide linkage. Clearly the DNA-immobilization reaction conditions did not increasethe yield by irreversibly denaturing the DNA, i.e., and thereby expose single-stranded deoxycytidylates. Inaddition, at the admittedly limited level of detection afforded by agarose gel electrophoresis, the restrictionenzyme digest results gave no evidence that the DNA-immobilization reaction conditions resulted in significantdeamination of deoxycytidylates or depurination (which would have affected the ApaL I results).The relative EtdBr-staining intensities of the two fragments from the Ssp I digests performed using freeand immobilized DNA were particularly informative about the nature of the deoxycytidylate-acyl hydrazidelinkage (Fig. 24, lanes 5-8). The larger (-.2100-bp) fragment was severely underrepresented in the digestsupernate, presumably because the overwhelming majority of these digestion products remained attached to thesupport. It therefore appears that the site of immobilization was a deoxycytidylate that could have been locatedanywhere along the 2,686 bp of linearized pUC 19 DNA (although it is possible that deoxycytidylates located inhot spots for transient helix-opening events were particularly reactive). The probability of this event havingoccurred somewhere along the length of the 2100-bp Ssp I-Kpn I fragment was therefore much higher than theprobability of the event having occurred somewhere along the length of the —600-bp Ssp I-Kpn I fragment. Therestriction enzyme digest results of Fig. 24 are therefore in good agreement with the results of Table 12, which139showed that the site of immobilization was not restricted to deoxycytidylates located in the single-strandedcaudemers of the DNA fragment. Hence as a first approximation (i.e., except for the extreme cases of very longor very short digestion products) the use of immobilized DNA as a restriction enzyme substrate results inrecoverable digestion products whose EtdBr-staining intensities are averaged—shorter digestion productsbecause of the nature of EtdBr staining, and longer digestion products by their greater probability of remainingon the solid support.140CHAPTER 2.4. DISCUSSIONA. THE IMMOBILIZATION OF DNA ONACYL HYDRAZIDE-DERIVATIZED SOLID SUPPORTS.1. Effective-concentration and catalytic effects.The transamination of cytosine moieties by acyl hydrazides is specific for single-stranded regions ofpolynucleotides (Hayatsu, 1976a, 1976b; Hayatsu & Ukita, 1966). This study showed that, at least when bufferTC was used, the transamination reaction was not efficient enough to immobilize significant amounts of DNAvia the terminal deoxycytidylates (3’ or 5’) of single-stranded caudemers of linear DNA restriction fragments.Some type of transient helix-opening event was therefore a prerequisite for the immobilization of significantamounts of double-stranded DNA on acyl hydrazide-derivatized solid supports. Hence one role of the couplingbuffer was to increase the effective concentration of reactive (i.e., single-stranded) deoxycytidylates, but withoutirreversibly denaturing the DNA (Zimmer & Venner, 1966; Zimmer et al., 1968). Another role of the couplingbuffer was to increase the rate that these transiently single-stranded deoxycytidylates were transaminated by acylhydrazides.The effective concentration of sterically accessible acyl hydrazides was also a factor affecting the yieldof immobilized DNA. Just as the nominal concentration of acyl hydrazides varies among different resins, so toodoes the fraction of these functional groups that are sterically accessible to DNA—e.g., as a result of thechemical nature, bead size, pore size, acyl hydrazide derivatization chemistry (there are several synthetic routes),and spacer arm (if any) of the matrix. Of the two resins examined in this study, Affi-Gel HZ appeared to have ahigher effective concentrationt of acyl hydrazides than Affi-Prep HZ. This was not surprising given the greaterporosity of Affi-Gel HZ, and it probably explains the greater yields of immobilized DNA obtained using thisresin (Tables 9 and 12). In this respect it should be noted that when Affi-Prep HZ was used at 30 °C (thetemperature used for the Affi-Gel HZ reactions), the yield of immobilized DNA was much less than that of thesame reaction at 37 °C (data not shown).t The nominal acyl hydrazide concentrations of these resins are not specified by the vendor, but both aredescribed in Bio-Rad product literature as having a binding capacity of 1-10 mg of periodate-oxidized IgG permL of resin.141The immobilization of double-stranded DNA on acyl hydrazide-derivatized solid supports is a mixed-phase reaction. The rheological properties of the resin suspension increased the efficiency of mixing, which inturn increased the effective concentration of accessible acyl hydrazides. The PEG 8000 and supplementary KC1in buffer TC minimized nonspecific adsorption and prevented clumping of the resin during the reaction, and thisprobably accounted for their beneficial effects on the yield of immobilized DNA. The rheological effect of thesupplementary KC1 was nontrivial, as the presence of 50 in)’l KC1 in the reaction must have significantlyreduced the effective concentration of single-stranded deoxycytidylates in the double-stranded DNA.In the series of experiments that were used to formulate the coupling buffer TC the nominalconcentrations of the reactants—deoxycytidylates and acyl hydrazides—were kept constant. However, theeffective concentrations of these reactants did not remain constant, but were instead highly dependent on thecoupling buffer used in the immobilization reaction. Distinguishing between effective-concentration andcatalytic effects is necessary for a proper understanding of the roles played by individual components of thecoupling buffer. Furthermore, regardless of what buffer is used in the DNA-immobilization reaction, theeffective concentration of acyl hydrazides on a particular solid support—i.e., acyl hydrazides that are stericallyaccessible to DNA—may be significantly less than the nominal concentration of this functional group on theresin.2. The roles played by individual components of the coupling buffer.a. The role of the hydrogen-ion buffer.Moderately acidic conditions are essential for the reaction of cytosine derivatives with acyl hydrazides(Gal-Or et al., 1967; Hayatsu & Ukita, 1964; Hayatsu et al., 1966; Kikugawa et al., 1967a). The mechanism forthis reaction has not been studied, but it probably shares several features of the bisulfite-catalyzedtransamination reactions of cytosine derivatives and amines (see Hayatsu, 1976a), and the buffer-catalyzeddeamination reactions of cytosine derivatives (Shapiro & Klein, 1966). Protonation of cytosine occurs undermoderately acidic conditions, which accentuates the electron-deficiency of the pyrimidine ring. By analogy withthe postulated mechanisms of deammation and amine transamination reactions, the electron-deficiency of the142protonated cytosine ring facilitates addition reactions across the 5,6 double bond, and saturation of the5,6 double bond in turn activates the pyrimidine ring for transamination.The primary catalytic effect of the hydrogen-ion buffer was due to its ability to maintain the appropriatehydrogen-ion concentration. However, by analogy with the addition-elimination reaction mechanismsmentioned above, two secondary catalytic effects may have also been exerted by the hydrogen-ion buffer.During the “addition phase” of this mechanism the carboxylic acid buffer may have acted catalytically by addingacross the 5,6 double bond (Seclor et al., 1974; Shapiro, 1981). The acid-catalyzed addition of carboxylic acidsacross carbon-carbon double bonds is a well-known reaction in simple olefins (see March, 1985). The anion ofthe carboxylic acid buffer (or its countenon) may have subsequently exerted another catalytic effect byincreasing the rate of abstraction of the hydrogen atom from C-5 during the “elimination phase” of the overallreaction mechanism (Sedor et al., 1974; Shapiro, 1981).The combination of these two secondary catalytic effects may explain why 4-aminobutanoic acid wassuch an effective hydrogen-ion buffer for the immobilization reaction. After the carboxylic acid group of thisbuffer added across the 5,6 double bond (and catalyzed the transamination reaction at C-4), the y-amino groupmay have subsequently catalyzed the elimination reaction intramolecularly, acting in the same fashion as theamino group of cysteamine (or the amino or hydroxyl group in related compounds such as cysteine or2-mercaptoethanol) during the catalysis of hydrogen-deuterium exchange at C-S in uridine (Hayatsu, 1976a;Heller, 1968; Kawai & Hillcoat, 1974; Wataya et al., 1973). The differing abilities of various hydrogen-ionbuffers to add across the 5,6 double bond and subsequently abstract the hydrogen atom from C-S may explainthe surprising variation in the yields of immobilized DNA obtained using the potassium salts of varioushydrogen-ion buffers (Table 5).Although the catalytic effects of the hydrogen-ion buffer were important, the pH of the immobilizationreaction must have also had a profound influence on the structure of the double-stranded DNA (Banville et al.,1986; Courtois et al., 1968; Gehring et a!., 1993; O’Connor et a!., 1981; Robinson et a!., 1992; Smol’janinovaet a!., 1982; Zimmer & Venner, 1966; Zimmer et a!., 1968). The effective concentration of reactive,single-stranded deoxycytidylates was greatly increased by the moderately acidic conditions of the reaction. This143phenomenon appeared to involve reversible, transient helix-opening events that could be completely suppressedby higher ionic-strength conditions.b. The role of tricarballylic acid counterion.After TCBA was selected as the hydrogen-ion buffer, various bases were examined for their influenceon the yield of immobilized DNA. Each base supplied one of three different kinds of countenon: an alkali metalion (Na+ or K+), or the conjugate acid of a nitrogen-containing base whose predominant form at pH 4.20 haseither one {NH4, TRIS-Ht BIS-TRTS-H, and DABCO-H) or two { (piperazine-2H)2and (cyclam.2H)2positive charges. There were significant differences in the yield of immobilized DNA when these bases wereused, differences that were attributed to catalytic or effective-concentration effects.The different results obtained with the alkali-metal-ion-containing buffers were presumably due todifferences in the effective concentration of deoxycytidylates in the immobilization reactions. However, it isvery difficult to explain why these cations would have significantly different effects on the structure of DNA atpH 4.20. It is also difficult to explain the superior performance of the buffers that contained piperazine orcyclam (TP and TC) on this basis. Furthermore, the “iT’ buffers severely inhibited the DNA-immobilizationreaction (Table 10). In these buffers the increased concentration of (piperazine2H)2+or (cyclam2H)2+probably caused a drastic reduction in the effective concentration of reactive single-stranded deoxycytidylates bystabilizing the double-stranded structure of the DNA.Piperazine and cyclam may have acted as better “elimination’ catalysts for the abstraction of thehydrogen atom from C-5 of the cytosine ring, and this may have accounted for the superior performance of thecoupling buffers that contained these bases (Sedor et al., 1974; Shapiro, 1981). Nitrogen-containing bases canpresumably act at this step, as the buffer-catalyzed deamination of cytidine proceeds at pH 5.0 in the presence ofpyridine-HC1, a noncarboxylate buffer (Shapiro & Klein, 1966). The protonated secondary amines of linearpolyamines have a high affmity for nucleic acids, and linear polyamines are able to migrate along DNAextremely rapidly (Besley et al., 1990; Frydman et at., 1992). All of the nitrogen atoms in piperazine andcyclam are secondary amine nitrogens, and hence the dications of these bases probably have a very high affinityfor DNA. At very low concentrations the structure-stabilizing effect of these dications on the DNA double-helix144was presumably more than offset by their catalytic effect during the elimination step of the reaction mechanism,whereas at higher concentrations (i.e., in the “TT” buffers; Table 10) the structure-stabilizing effect of(piperazine-2H)2or (cyclam-2H)2predominated.c. The role ofpolyetliylene glycol.DNA undergoes a polymer-and-salt-induced (N,) transition from an extended coil to a condensed“globule” form in solutions containing appropriate concentrations of PEG and salt (Lerman, 1971, 1973; Sikorav& Church, 1991). The N’ transition is a reversible phenomenon that does not result in the denaturation ofdouble-stranded DNA. However, depending on the properties of the DNA and the solvent, f-condensation maycause aggregation of DNA into a rapidly sedimentable form (Lerman, 1971, 1973).Given that the f-condensation of DNA increases the rate of DNA renaturation (Amasino, 1986;Silcorav & Church, 1991), and would therefore decrease the effective concentration of reactive single-strandeddeoxycytidylates, it is probably unlikely that f-condensation conditions are compatible with the immobilizationof double-stranded DNA on acyl hydrazide-derivatized solid supports. A N,-transition may have been the reasonfor the precipitous drop in the yield of immobilized DNA when the supplementary KC1 concentration was raisedfrom 100 to 200 mM (Table 8). When the TCBA concentration was doubled from 8 to 16 mM in the “T” and“TI”’ buffers, the corresponding increase in the concentration of the polyamine dication { (piperazine2H)2+or(cyclam-2H)2jmay also have provoked a f-transition (Table 10).As mentioned earlier, the PEG 8000 and supplementary KC1 in buffer TC probably increased theefficiency of mixing of the resin during the immobilization reaction, and this probably accounted for thebeneficial effects of these solutes on the yield of immobilized DNA. The effect of macromolecular crowdingwas probably minor, especially for lower Mr DNA, as the (fmal) concentration of PEG 8000 could be lowered to2% w/v with little effect on the immobilization of the 51-bp duplex HSE probes (data not shown).Finally, it should be noted that PEG 8000 cannot be used with certain solid supports—one noteworthyexample being very porous, acyl hydrazide-derivatized polyacrylamide-based resins. Whether or not to includethis solute in the coupling buffer, and at what concentration, should therefore be determined empirically,145depending on the type of solid support and the size and concentration of the DNA to be used in theimmobilization reaction.3. Important unanswered questions about the immobilization chemistry.a. The distribution and average number ofattachment points per DNA fragment.Two related questions concerning the immobilization of unmodified, double-stranded DNA on acylhydrazide-derivatized solid supports are the distribution and average number of attachment points per DNAfragment. The answers to these questions will probably vary depending on the length of the DNA fragment, andthe presence of sequences within it that may act as hot spots for immobilization due to their propensity fortransient helix-opening events. The duration of the immobilization reaction and the nominal concentration,steric accessibility, and distribution of acyl hydrazide groups on the resin may also help to determine thedistribution and average number of attachment points per DNA fragment.These two questions were not even answered for the specific reaction conditions, resins, and DNA usedin this study. However, the results of the microscale affinity chromatography of the putative ceHSBFm activityperformed using poly(hse-A•hse-T) immobilized on Affi-Gel HZ, and the restriction enzyme digests performedusing linearized pUC 19 DNA immobilized on Affi -Prep HZ, indicated that most DNA fragments were probablyimmobilized by a single attachment point (Figs. 23 and 24).b. The deamination and depurination ofDNA at pH 4.2.It is pointless to use acyl hydrazide-derivatized solid supports for the immobilization of DNA if themoderately acidic conditions required by the reaction chemistry result in a high frequency of mutations. AtpH 4.2 the most frequent types of mutations that can occur are the deamination of cytosine, adenine and guaninemoieties, and the hydrolysis of the N-glycosidic bond of all four deoxyribonucleotides (Lindahi, 1993; Lindahl& Nyberg, 1974; Shapiro & Danzig, 1972). At pH 4.2 the deamination of cytosine is much more significantthan the deamination of purines, and the hydrolysis of the N-glycosidic bond of purine residues is much moresignificant than depyrimidination (Lindahl, 1993). In the following discussion it should also be noted that theN-glycosidic bond of a ribonucleotide is much more resistant to hydrolysis than that of the corresponding146deoxyribonucleotide (Lindahi, 1993). Furthermore, although the rate of hydrolysis of the N-glycosidic bond ofdeoxyribonucleotides in single-stranded DNA is only about four times the corresponding rate of hydrolysis indouble-stranded DNA, the rate of deamination of cytosine moieties in single-stranded DNA is about 100-200times the corresponding rate of hydrolysis in double-stranded DNA (Lindahi, 1993; Lindahi & Nyberg, 1974;Shapiro & Danzig, 1972).This study did not measure the extent of deamination of cytosine moieties or the extent of depurination,and it is not clear which mutation would be more frequent under the immobilization reaction conditions used inthis study. However, if depurination was less frequent than deamination then it is unlikely that either event wasvery significant. This is because the competing deamination reaction is much less efficient than thetransamination of cytosine derivatives by acyl hydrazides. As an example, when Gal-Or et al. (1967) incubatedlabeled yeast RNA at 37 °C in 2 M malonyl hydrazide, pH 4.2, for 90 hours, and then analyzed the hydrolyzedRNA to determine its ribonucleotide composition, they reported that the levels of AMP, GMP, and UMP wereunchanged within experimental error. In addition, when Hayatsu & Ukita (1964) incubated adenosine,guanosine, uridine, and cytidine in separate reactions at 37 °C with 1 M semicarbazide, pH 4.2, for 125 hours,only cytidine was found to react, and the only product observed was the 4-deamino-4-semicarbazino cytidinederivative. The absence of any significant deamination of cytosine moieties after extended incubation at pH 4.2and 37 °C was confirmed in subsequent studies using nucleosides and nucleotides (Hayatsu et al., 1966) andRNA (Hayatsu & Ukita, 1966).Unfortunately, the above results provide little useful information about the potential significance ofdepurination under the reaction conditions used in this thesis, as these studies used RNA, not DNA. However,when Reisfeld et al. (1987) attached biotin hydrazide to DNA probes via the bisulfite-catalyzed transaminationof cytosine moieties (in a 24-hour reaction at pH 4.5 using 1 M bisulfite), the hybridization properties of theprobes were unaffected, suggesting that there was no significant depurination of the DNA (or deamination ofcytosine moieties) during the course of the transamination reaction. Finally, the results of the microscale affmitychromatography of the putative ceHSBFm activity performed using poly(hse-A.hse-T) immobilized onAffi-Gel HZ, and the restriction enzyme digests performed using linearized pUC 19 DNA immobilized on147Affi-Prep HZ, also indicated that under the standard immobilization reaction conditions used in this thesis (6 h,30-37 °C, using buffer TC at pH 4.20) the extent of deamination or depurination was probably not verysignificant.c. The possibility ofacyl hydrazone formation at apurinic sites in DNA.Depurination of DNA yields apurinic (AP) sites. The sugar at an AP site is an aldose, and the cyclichemiacetal form of this sugar exists in equilibrium with the acyclic aldehyde form. Opening of the2-deoxyribofuranose ring at an AP site would therefore yield a potentially reactive aldehyde, and under theconditions used for immobilizing DNA in this study, this aldehyde might react with an acyl hydrazide on thesolid support to yield an acyl hydrazone (Fig. 25). Although this alternative reaction chemistry was neverinvestigated systematically in this study, the relevant data in the literature, as well as the evidence of Fig. 22 andTables 8-10, all argue against the “acyl hydrazone” hypothesis outlined in Fig. 25.The first problem encountered by the acyl hydrazone hypothesis is the extent of depurination of theDNA during a 6-hour incubation at pH 4.2 and 37 °C. For the purposes of discussion, one can simply treat all ofthe linearized double-stranded pUC 19 DNA in the immobilization reaction as single-stranded DNA, andestimate an upper limit on the extent of depurination during the reaction. This way the ionic strength of thereaction can be ignored, as a worst-case scenario is being considered (if the extent of depurination ofdouble-stranded DNA was being estimated, the ionic strength of the reaction could not be ignored).The data of Lindahl & Nyberg (1972) show that (i) the Arrhenius plots for the depurination ofdouble-stranded and single-stranded DNA samples have the same slope, and that this slope is unchanged whenmeasured at pH 5.0 and pH 6.0; and (ii) at 70 °C the difference in the value of the log of the rate constants forthe depurination of double-stranded DNA at pH 7.4 and pH 4.2 is 2.7 (a temperature of 70 °C was required toobtain a measurable rate of depunnation). Thus at 37 °C, the rate of depurination of single-stranded DNA atpH 4.2 is probably about 500 timest as fast as the rate of depurination at pH 7.4.t With an acid-catalyzed reaction such as depurination one would expect that the rate would increase 10-foldwith each decrease of one pH unit. However, above pH 6.0, depurination may also proceed by apH-independent pathway, and the observed rate of depurination at pH 7.4 is five times greater than what wouldbe expected solely on the basis of the acid-catalyzed reaction (Lindahl & Nyberg, 1972).148rinoring opening00purine0-000-013-eliminationstrand scissionH2NN%%__ resinacyl hydrazoneformationresin®—cFigure 25. A hypothetical mechanism for DNA immobilization via acyl hydrazone formation.149Shapiro (1981) has estimated that at 37 °C and pH 7.4, one depurination event occurs in 2 x 106deoxyribonucleotide residues of single-stranded DNA every 2.5 hours; hence —2.4 depurination events occur in2 x 106 deoxynucleotide residues of single-stranded DNA every 6 hours. Thus one can estimate that 1,200depurination events occur in 2 x 106 deoxynucleotide residues of single-stranded DNA every 6 hours at 37 °Cand pH 4.2. Another way of expressing this value is that —1 depurination event occurs per 1,700deoxynucleotide residues of single-stranded DNA every 6 hours at 37 °C and pH 4.2.Thus even if all of the linearized double-stranded pUC 19 DNA in the immobilization reaction istreated as single-stranded DNA, it is difficult to accept that the extent of depurination would be adequate toprovide enough potentially reactive sites for any detectable immobilization to occur via acyl hydrazoneformation. Each double-stranded molecule of pUC 19 (or 2 x 2,686 = 5,372 deoxyribonucleotides) would onlyhave a maximum of -.3 potentially reactive AP sites. This number is only slightly greater than the number ofterminal (single-stranded) deoxycytidylates (two) on each linearized molecule of pUC 19 used in theimmobilization reaction.There is another serious problem for the acyl hydrazone hypothesis: the reaction of aldoses withcarbonyl reagents involves attack of the free aldehyde, not the cyclic hemiacetal form of the sugar (reviewed byCapon, 1969). Indeed, even semicarbazide reacts by attacking the free aldehyde form of aldoses (to form thecorresponding semicarbazones), and semicarbazide is a very reactive acyl hydrazide (Capon, 1969). Yet the(cyclic) 2 -deoxyribofuranose at an AP site is the predominant form of this sugar in solution, and although thepresence of acid (or base) catalyzes the rapid interconversion of the two isomers, only 1% of the sugar is in thereactive aldehyde form in aqueous solution (reviewed in Lindahi, 1993). In contrast, when glycoproteins orRNA are treated with periodate, this reagent cleaves vicinal diols to form dialdehydes. These dialdehydes areincapable of cyclization, and hence are fully available to react with functional groups on an acylhydrazide-derivatized resin to form acyl hydrazones.Furthermore, instead of reacting with an acyl hydrazide, the reactive aldehyde form of 2’-deoxyribose atan AP site would probably undergo successive 13-elimination reactions, resulting in a degraded sugar and strandscission (Shapiro, 1981; Lindahi, 1993). Yet the restriction enzyme digestion results of Fig. 24 (obtained using150DNA that had been immobilized on Affi-Prep HZ) showed the expected digestion products, and did not showany degradation of the DNA.Finally, the immobilization of double-stranded DNA on acyl hydrazide-derivatized solid supports isstrongly catalyzed by bisulfite (Fig. 22). In the experiment whose results are described in Fig. 22, one reactionwas performed without bisulfite, and here essentially no DNA was immobilized on the resin (lane 2). This lackof reactivity was probably because the pH and the ionic strength of the uncatalyzed transamination reaction weretoo high. However, the results of lanes 3 and 4 in Fig. 22 show that the addition of bisulfite to the reactionresulted in a dramatic increase in the efficiency of immobilization. The only plausible mechanism for thisimmobilization reaction is the bisulfite-catalyzed transamination of deoxycytidylates by acyl hydrazides. Theseresults are entirely consistent with the results of Reisfeld et al. (1987), who attached biotin hydrazide to DNAprobes via the bisulfite-catalyzed transamination of cytosine moieties (in a 24-hour reaction at pH 4.5 using 1 Mbisulfite). Bisulfite would be totally unnecessary and in fact inhibitory if (nominally) double-stranded DNA wasimmobilized via acyl hydrazone formation at AP sites (sodium bisulfite would be inhibitory due to a reduction inthe rate of depurination of double-stranded DNA with increasing ionic strength).Oxidized glycoproteins can be immobilized on acyl hydrazide-derivatized resins (i.e., via acylhydrazone formation) in buffers of quite significant ionic strength (e.g., 0.1 M NaOAc, 0.15 M NaCl, pH 5.5),viz., the acyl hydrazone reaction itself is relatively insensitive to salt concentration in this range (O’Shannessy &Hoffman, 1987). Therefore if acyl hydrazone formation did account for the immobilization of the DNA in thisstudy, the dramatic differences in the results shown in Table 10 (cf. the ‘T” and “IT” buffers), as well thoseshown in Tables 8 and 9 (cf. the results of the 16-hour and 4-hour reactions), could only be attributed todifferences in the rate of depurination. Yet this effect is only modest—there is only a fourfold difference in therates of depurination of double-stranded and single-stranded DNA (Lindahl, 1993)—whereas the effect of salton the DNA immobilization reaction studied herein was much more dramatic. No DNA was immobilized in a16-hour reaction at pH 4.20 if 200 mM KC1 was present (Table 8), whereas in an otherwise identical reaction,0.61 mg of DNA per mL of resin could be immobilized in 4 hours if 50 mM KC1 was present (Table 9).151B. APPLICATIONS USING DNA IMMOBILIZED ONACYL HYDRAZIDE-DERIVATIZED SOLID SUPPORTS.1. Some properties of acyl hydrazide-derivatized solid supports.a. Versatility, convenience, and safety.Versatility, convenience, and safety are three important attributes of commercially available acylhydrazide-derivatized solid supports. This derivatization chemistry can be used with various types of parentsupports, including agarose- and polyacrylamide-based resins (Inman, 1974; O’Shannessy & Wilchek, 1990;Wilchek et al., 1984). Many of these resins are available commercially, as acyl hydrazides are relatively stableto prolonged storage in an appropriate solvent. The use of a commercially available, acyl hydrazide-derivatizedsolid support for the affinity chromatography of sequence-specific DNA-binding proteins would spare theresearcher the hazards associated with using CNBr, and the expense and regulatory difficulties associated withthe purchase, shipping, storage, and proper disposal of this extremely toxic reagent.b. DNA-binding capacities ofAffi-Prep HZ and Affi-Gel HZThe DNA-binding capacities of the commercially available, acyl hydrazide-derivatized solid supportsused in this study were greater than those of CNBr-activated resins, but lower than those achieved using severalother resins, e.g., epoxide activated latex particles (Inomata et al., 1992), (strept)avidin-derivatized resins (usedin conjunction with biotinylated deoxyribonucleotides, e.g., Gabrielsen et al., 1989), or N-hydroxysuccinimidedenvatized resins (used in conjunction with deoxyribonucleotides bearing tethered nucleophiles, e.g., Larson &Verdine, 1992).c. “Uncapped” or unreacted acyl liydrazides.In both of the applications that were described in this study—DNA-affinity chromatography andrestriction enzyme digests—the unreacted acyl hydrazides on the solid supports were not “capped” after theDNA-immobilization reaction. One form of capping that might be contemplated would be the formation of asimple acyl hydrazone (or possibly acylation); however, the vendor’s instructions for the use of Affi-Prep HZand Affi-Gel HZ do not mention and hence presumably do not recommend that unreacted acyl hydrazides on152these resins be capped. Nevertheless, it may still be advisable to avoid exposing an acyl hydrazide-derivatizedresin (prepared as described in this study) to oxidized carbohydrates (particularly under acidic conditions), as the“uncapped” acyl hydrazide groups may react with simple aldehydes (or ketones) on such sugars to form acylhydrazones. Un-oxidized carbohydrates would probably not react with acyl hydrazide groups because (i) thereis no free aldehyde available in a nonreducing sugar (the anomeric carbons are all involved in glycosidiclinkages); and (ii) in a reducing sugar the carbonyl of an acyclic aldose or ketose would be much more prone toreact intramolecularly to form the unreactive, cyclic hemiacetal or hemiketal form of the sugar, respectively.Many eukaryotic transcription factors and nuclear proteins are glycosylated (Jackson & Tjian, 1988,1989; Kan & da Silva, 1986; Lichtsteiner & Schibler, 1989), including the only RNA polymerase II regulatoryfactor from C. elegans that has been studied biochemically (Okai, 1986a). Oxidation of the sugar residues onthese proteins is potentially a problem, as these residues may then react with ‘uncapped” acyl hydrazides on theresin. However, the concentration of proteins bearing oxidized sugar residues is usually so low that it does notpresent a problem in chromatographic separations that use acyl hydrazide-derivatized resins.In this study uncapped” (unreacted) acyl hydrazides did not appear be a problem. Furthermore, theseresidues are essentially uncharged above pH 4.5 (Lindegren & Niemann, 1949), which means that under mostoperating conditions these residues do not confer any undesirable ion-exchange properties on the resin. This isnot the case with unreacted cyanate esters, which after capping with primary amines yield isourea derivatives(Wilchek et al., 1984). Under most operating conditions these isourea linkages will be positively charged, andhence may confer some undesirable anion-exchange properties on CNBr-activated resins (Wilchek et a!., 1984).This problem is of particular concern given the extremely high degree of activation that is required to prepareCNBr-activated resins for the immobilization of double-stranded DNA.d. The rate of loss of immobilized DNA.The rate of loss of immobilized DNA from acyl hydrazide-derivatized solid supports is probably themost serious problem affecting this type of resin. The rate of loss of poly(dI-dC).poly(dI-dC) from acylhydrazide-derivatized solid supports is so high that such resins are probably unsuitable for routine use (e.g., as aprecolumn for removing nonspecific DNA-binding proteins). One significant advantage of CNBr activation for153the immobilization of double-stranded DNA is the stability of the resulting linkage (Arndt-Jovin et al., 1975; butsee also Bünemann et a!., 1982). However, the rate of loss of pUC 19 DNA from acyl hydrazide-derivatizedsolid supports (30-40% after 10 months at 4 °C; Table 14) should be kept in perspective. Even after 10 monthsstorage at 4 °C, the amount of pUC 19 DNA remaining (600-770 j.iglmL) on either of the two acyl hydrazidederivatized solid supports that were examined in this study was still much greater than the amount of double-stranded DNA that can be immobilized using CNBr-activated agarose (—100 p.g of DNA per mL of resin;Kadonaga, 1991). In most cases the rate of loss of DNA from acyl hydrazide-derivatized solid supports willprobably not adversely affect a limited-duration experiment such as an affinity chromatography separation.In the experiments to determine the loss of DNA from both types of resin (Table 14), no antioxidantswere added to the storage buffer because of the nature of the assay that was to be used later to quantitate theselosses. In addition, the extent of these losses may have been affected by the composition of the storage buffer.It may therefore be worthwhile to examine more systematically the composition of the storage buffer—includingthe possible inclusion of antioxidants—in an attempt to reduce the losses of DNA from these resins.2. Immobilized DNA as an affinity chromatography ligand.The results of the microscale affinity chromatography of the putative ceHSBFm activity showed thatpoly(hse-A.hse-T) immobilized on Affi-Gel HZ was able to function as a sterically accessible DNA-affinitychromatography ligand. Ligation of the DNA prior to immobilization was essential, as an initial attempt to usethe 5 1-bp hse-A•hse-T monomer as an affinity ligand was unsuccessful. In retrospect, the failure of this attemptwas understandable given the nature of the immobilization reaction. The average size of the DNA concatemersproduced by the ligation reaction will not only affect the yield of immobilized DNA, it will also determine howaccessible that DNA will be and hence how well that DNA will function as an affinity chromatography ligand.The unsuccessful attempt to use the 51-bp hse-A•hse-T monomer as an affinity ligand indicated that theputative ceHSBFm activity did not have any affinity for the acyl hydrazide matrix itself. However, the affinitychromatography experiment whose results were reported in Fig. 23 should have included an appropriate controlto show that this observation (lack of binding to the parent support) was reproducible.154Given enough biological starting material and the requisite oligodeoxynbonucleotides, other controlexperiments could also have been performed. In many cases one can apply the sample to an agarose-basedcolumn containing immobilized nonspecific DNA (ideally, concatemerized DNA that contains sequencesidentical to the flanking sequences on the DNA used for the sequence-specific affinity column), and recover thesequence-specific DNA-binding protein of interest in the flow-through fraction. The immobilized nonspecificDNA retains nonspecific DNA-binding proteins, and the column itself also retains any proteins that bind to theagarose-based support. In many cases nonspecific competitor DNA (—1,000 bp in length) is also added prior toloading to prevent nonspecific DNA-binding proteins from being retained by a sequence-specific DNA-affinitycolumn (Kadonaga, 1991; Kadonaga & Tjian, 1986), and in many cases this step is viewed as crucial to thesuccessful purification of sequence-specific DNA-binding proteins. However, the complete removal ofnonspecific DNA-binding proteins using these approaches is rarely achieved, and exploratory experiments arerequired to determine the amount of nonspecific DNA (on a solid support or in the mobile phase) necessary toremove the bulk of these proteins. Furthermore, in some cases the recovery of the desired protein in theflow-through from a nonspecific-DNA-agarose column, and the addition of nonspecific DNA prior to theloading of a sequence-specific DNA-affinity column, are not possible (e.g., Kaufman et al., 1989).Other control experiments ensure that a sequence-specific DNA-affinity column selectively retains aDNA-binding protein that is specific for the “target’ sequence (cognate binding site) present in multiple copieson the concatemerized DNA used to prepare the column, and not a DNA-binding protein that is specific forsome other (i.e., flanking) sequences that are also present in multiple copies on the DNA ligand. Theseexperiments involve the sequential use of two sequence-specific DNA-affinity columns, both of which areprepared with concatemerized DNA that contains multiple copies of the cognate binding site, but which haveunrelated flanking DNA sequences (Kadonaga, 1991; Kadonaga & Tjian, 1986). The sequence-specificDNA-binding protein of interest should bind to both columns.As was stated earlier in the Results Section, the primary objective of the experiment whose results werereported in Fig. 23 was to determine if poly(hse-A•hse-T immobilized on Affi-Gel HZ was able to function as asterically accessible DNA-affinity chromatography ligand. Unfortunately there were severe limitations on the155amount of nematode eggs that I was able to prepare for use in the experiments described in this thesis, and forreasons that are beyond the scope of this discussion I had to assume the entire financial burden associated withthe purchase of the oligodeoxyribonucleotides used in this work. It is regrettable that the experiment whoseresults were reported in Fig. 23 was not repeated altogether—with the appropriate controls—to ensure that theseresults were reproducible. However, even in the absence of these controls, the available evidence still suggeststhat poly(hse-A.hse-T) immobilized on Affi-Gel HZ was able to function as a sterically accessible DNA-affinitychromatography ligand and retain a significant fraction of the putative ceHSBF activity present in a crudenuclear extract applied to the column.One advantage of acyl hydrazides is that the derivatization chemistry for their synthesis is compatiblewith a variety of parent supports. Unfortunately, I was unable to examine the suitability of Affi-Prep HZ as asolid support for the affinity chromatography of sequence-specific DNA-binding proteins. Although almost allsuch experiments have been carried out with agarose-based resins (because of their porosity and the use of CNBractivation for the immobilization of DNA), agarose-based resins may not be ideal for this application. Histoneshave been reported to bind irreversibly to Sepharose (see Svaren et al., 1987), which suggests that these proteinsalso bind irreversibly to other agarose-based resins. Other types of solid supports (e.g., Affi-Prep HZ) may notshare this potentially troublesome property, and for this and other reasons these supports (or other syntheticpolymer-based resins) may be more suitable for the preparation of DNA-affinity chromatography matrices.3. Immobilized DNA as a substrate for enzyme-catalyzed reactions.The utility of Affi-Prep HZ for the immobilization of unmodified double-stranded DNA and itssubsequent use as a solid-phase substrate for enzymatic reactions is particularly significant. The parent support,a macroporous, poly(methyl methacrylate)-based HPLC resin, tolerates elevated temperatures and high flowrates. This malces the support suitable for a variety of applications in recombinant DNA technology,applications which could easily assume much more importance than the use of these resins for sequence-specificDNA-affinity chromatography. One of the most useful attributes of immobilized DNA as a substrate forenzymatic manipulations is the ability to carry out such reactions automatically using robotic workstations.156Two important parameters affecting the utility of immobilized DNA as a substrate for enzymaticreactions are the distribution and average number of attachment points per DNA fragment. As was pointed outearlier, many factors will determine the values of these parameters. In many experiments it would be desirableto have randomly distributed, single-point attachment of the DNA fragments to the resin. 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Biochemistry and Molecular Biology
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Some new approaches to conventional methods for the study of protein-DNA interactions
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