Medicine, Faculty of
Biochemistry and Molecular Biology, Department of
DSpace
UBCV
Behar, Hila
Doctor of Philosophy - PhD
University of British Columbia
A defining feature of plants, the plant cell wall is an essential structure that provides mechanical support, cell shape, and protection from pathogens and the environment. Cell walls are dynamic structures that are composed of a network of crystalline cellulose microfibrils, amorphous polysaccharides, and structural proteins. The synthesis, maintenance, and modification of the plant cell wall polysaccharides are carried out by carbohydrate active-enzymes (CAZymes). Xyloglucan endotransglycosylase/hydrolase (XTH) are secreted cell wall enzymes that restructure the cell wall matrix glycan, xyloglucan, thereby facilitating cell wall remodelling and growth. The plant endo-glucanase 16 (EG16) enzymes were recently identified as a unique clade that is structurally and functionally intermediate between the bacterial mixed-linkage β(1,3)/(1,4)-glucanases and the plant XTHs in glycoside hydrolase family 16 (GH16). Correspondingly, EG16 members were broadly specific towards the cell wall polysaccharides mixed-linkage β-glucan (MLG) and xyloglucan, as well as soluble cellulosic substrates. Although EG16 activity and protein structure have been demonstrated, their biological role is unknown. This thesis employs a multi-disciplinary approach to the characterization of the newly designated EG16 group. A comprehensive census of the publicly available genome and transcriptomes showed that EG16 orthologs were found in all plant clades spanning nearly 500 million years of evolution. Notably, the presence of EG16 orthologs observed in the charophyte green algae species that are the most phylogenetically distant from angiosperms, support that EG16-like genes are ancestors to extant EG16 and XTH genes. EG16 substrate specificities, primary hydrolysis site, and digest products are consistent between vastly different plant species. Elucidation of the biological role of EG16 in the moss Physcomitrella patens and a hybrid poplar species, two different distantly related land plants, show gene expression in young tissues and leaderless cell wall localization. In P. patens, EG16 deletion yielded larger plants that senesced earlier than wild-type, whereas, in poplar, downregulation did not affect growth. This thesis serves as a foundation for the EG16 clade and informs their evolutionary history, conserved activity, cell wall localization, and involvement in growth. The research opens the door to investigate unexplored topics such as the EG16 specific secretion mechanism and their role in MLG-containing cell walls.
UNDERSTANDING THE DIVERSITY, BIOCHEMISTRY, AND BIOLOGICAL ROLE OF ENDO-GLUCANASE 16 ENZYMES IN THE CONTEXT OF PLANT EVOLUTION by Hila Behar B.Sc., Tel Aviv University, 2014 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF Doctor of Philosophy in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Biochemistry and Molecular Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2021 © Hila Behar, 2021 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Understanding the diversity, biochemistry, and biological role of endo-glucanase 16 enzymes in the context of plant evolution submitted by Hila Behar in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biochemistry and Molecular Biology Examining Committee: Prof. Harry Brumer, Chemistry, UBC Supervisor Prof. A. Lacey Samuels, Botany, UBC Supervisory Committee Member Prof. Shawn D. Mansfield, Forestry, UBC Supervisory Committee Member Prof. Ljerka Kunst, Botany, UBC University Examiner Prof. Lawrence McIntosh, Biochemistry and Molecular Biology, UBC University Examiner Additional Supervisory Committee Members: Prof. Nobuhiko Tokuriki, Biochemistry and Molecular Biology, UBC Supervisory Committee Member iii Abstract A defining feature of plants, the plant cell wall is an essential structure that provides mechanical support, cell shape, and protection from pathogens and the environment. Cell walls are dynamic structures that are composed of a network of crystalline cellulose microfibrils, amorphous polysaccharides, and structural proteins. The synthesis, maintenance, and modification of the plant cell wall polysaccharides are carried out by carbohydrate active-enzymes (CAZymes). Xyloglucan endotransglycosylase/hydrolase (XTH) are secreted cell wall enzymes that restructure the cell wall matrix glycan, xyloglucan, thereby facilitating cell wall remodelling and growth. The plant endo-glucanase 16 (EG16) enzymes were recently identified as a unique clade that is structurally and functionally intermediate between the bacterial mixed-linkage β(1,3)/(1,4)-glucanases and the plant XTHs in glycoside hydrolase family 16 (GH16). Correspondingly, EG16 members were broadly specific towards the cell wall polysaccharides mixed-linkage β-glucan (MLG) and xyloglucan, as well as soluble cellulosic substrates. Although EG16 activity and protein structure have been demonstrated, their biological role is unknown. This thesis employs a multi-disciplinary approach to the characterization of the newly designated EG16 group. A comprehensive census of the publicly available genome and transcriptomes showed that EG16 orthologs were found in all plant clades spanning nearly 500 million years of evolution. Notably, the presence of EG16 orthologs observed in the charophyte green algae species that are the most phylogenetically distant from angiosperms, support that EG16-like genes are ancestors to extant EG16 and XTH genes. EG16 substrate specificities, primary hydrolysis site, and digest products are consistent between vastly different plant species. Elucidation of the biological role of EG16 in the moss Physcomitrella patens and a hybrid poplar species, two different distantly related iv land plants, show gene expression in young tissues and leaderless cell wall localization. In P. patens, EG16 deletion yielded larger plants that senesced earlier than wild-type, whereas, in poplar, downregulation did not affect growth. This thesis serves as a foundation for the EG16 clade and informs their evolutionary history, conserved activity, cell wall localization, and involvement in growth. The research opens the door to investigate unexplored topics such as the EG16 specific secretion mechanism and their role in MLG-containing cell walls. v Lay Summary Plant cell walls are essential structures that encompass the plant cells and are composed of mainly carbohydrate constituents. Cell wall carbohydrates are synthesized, modified, and maintained by enzymes. A new group of enzymes was recently discovered, called endo-glucanase 16 (EG16), which are evolutionarily related to a well-researched group of cell wall enzymes xyloglucan endotransglycosylase/hydrolase (XTH) that carry out cell wall rearrangements. While numerous copies of XTHs are found in all plant genomes, EG16s are only found once per genome and are missing from many flowering plants. In addition, EG16s are highly similar in vastly different plant species spanning 500 million years of evolution. Despite this, their biological role remains unknown. This thesis explores the diversity of EG16 enzymes across the plant lineages and investigates the biological role in two distantly related species, spreading earthmoss and hybrid poplar trees. vi Preface Chapter 1: Introduction. This chapter was written by the author (Hila Behar) and reviewed by Dr. Harry Brumer (Professor, Ph.D. supervisor), Dr. A. Lacey Samuels (Professor, Committee member), and Dr. Shawn D. Mansfield (Professor, Committee member). Chapter 2: Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG16 and XTH proteins in plant lineages. This chapter was co-written by the author (Hila Behar) and Dr. Harry Brumer (Professor, Ph.D. supervisor). The chapter was reviewed and edited by Dr. Sean Graham (Professor, Botany, UBC). Sequence extraction from public resources, in‐house database construction, bioinformatics analyses and figure assembly were done by the author. A version of this chapter was published (Behar et al., 2018), and a continuation of this analysis was subsequently carried out by the author and supervisor and integrated into another publication in the form of one figure ((Jiao et al., 2020); Figure 5B). Behar, H., Graham, S.W. and Brumer, H. (2018) Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG16 and XTH proteins in plant lineages. Plant J, 95, 1114-1128. Jiao, C., Sørensen, I., Sun, X., Sun, H., Behar, H., Alseekh, S., Philippe, G., Palacio Lopez, K., Sun, L., Reed, R., Jeon, S., Kiyonami, R., Zhang, S., Fernie, A.R., Brumer, H., Domozych, D.S., Fei, Z. and Rose, J.K.C. (2020) The Penium margaritaceum Genome: Hallmarks of the Origins of Land Plants. Cell, 181, 1097-1111.e1012. Chapter 3: Conservation of endo-glucanase 16 (EG16) activity across highly divergent plant lineages. This chapter was co-written by the author (Hila Behar) and Dr. Harry vii Brumer (Professor, Ph.D. supervisor), with contributions from Dr. Kazune Tamura (Brumer Lab, UBC), Dr. Daniel J. Cosgrove (Professor, Biology, Pennsylvania State University), and Edward R. Wagner (Biology, Pennsylvania State University). The author carried out sequence analysis, cloning, protein production and purification, kinetic analysis, and enzyme product analysis. Dr. Kazune Tamura (Brumer Lab, UBC) produced protein, prepared crystallography trials, performed homology modelling, and analyzed the resulting data. Edward R. Wagner produced protein and performed all experiments related to the biomechanical assays at Penn State with supervision by Prof. Daniel J. Cosgrove. A version of this chapter was submitted for initial review. Chapter 4: Physcomitrella patens endo-glucanase 16 is involved in the cell wall development of young tissue. This chapter was written by the author (Hila Behar) and revised by Dr. Harry Brumer (Professor, Ph.D. supervisor) and Dr. A. Lacey Samuels (Professor, Committee member). The author carried out all experimental work, data analysis and figure assembly. Chapter 5: Functional analysis of Populus endo-glucanase 16 (EG16). This chapter was written by the author (Hila Behar). The chapter was reviewed and edited by Dr. Harry Brumer (Professor, Ph.D. supervisor), Dr. A. Lacey Samuels (Professor, Committee member) and Dr. Shawn D. Mansfield (Professor, Committee member). The author performed genotyping, RT-qPCR analysis, expression analysis, plant growth phenotyping, and immunofluorescent microscopy. Yaseen Mottiar and Rohan Chandra (Mansfield Lab, UBC) performed the cloning and poplar transformation assays. Alegra Grappadelli (Heinrich Heine University Düsseldorf) performed MALDI-imaging and made the relevant figures. Prof. Markus Pauly (Heinrich Heine University Düsseldorf) supervised MALDI-imaging experiments. viii Chapter 6: Conclusions. This chapter was written by the author (Hila Behar) and reviewed by Dr. Harry Brumer (Professor, Ph.D. supervisor), Dr. A. Lacey Samuels (Professor, Committee member), and Dr. Shawn D. Mansfield (Professor, Committee member). ix Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ......................................................................................................................... ix List of Tables .............................................................................................................................. xvi List of Figures ............................................................................................................................ xvii List of Supplementary Material ............................................................................................... xxi List of Abbreviations ............................................................................................................... xxiii Acknowledgements .................................................................................................................. xxiii Dedication ...................................................................................................................................xxx Chapter 1: Introduction ................................................................................................................1 1.1 Plant cell walls ................................................................................................................ 1 1.1.1 Plant cell wall types .................................................................................................... 2 1.1.2 Plant cell wall polysaccharides ................................................................................... 5 1.1.2.1 Cellulose ............................................................................................................. 5 1.1.2.2 Pectin................................................................................................................... 5 1.1.2.3 Hemicellulose ..................................................................................................... 6 1.1.2.3.1 Mannans ........................................................................................................ 7 1.1.2.3.2 Xylans............................................................................................................ 9 1.1.2.3.3 Xyloglucan .................................................................................................... 9 1.1.2.3.4 Mixed-linkage β-glucan .............................................................................. 10 1.1.3 Cell wall proteins ...................................................................................................... 11 x 1.2 Carbohydrate-active enzymes (CAZymes) ................................................................... 12 1.2.1 Carbohydrate-Active enZymes (CAZy) database ..................................................... 12 1.2.2 Glycoside hydrolase family 16 (GH16) .................................................................... 12 1.2.2.1 Bacterial mixed-linkage glucanases (MLGases) .............................................. 13 1.2.2.2 Xyloglucan endotransglucosylase/hydrolase (XTH) enzymes ......................... 14 1.2.2.3 Endo-glucanase 16 (EG16) enzymes ................................................................ 15 1.3 Model organisms ........................................................................................................... 20 1.3.1 Plant model systems .................................................................................................. 21 1.3.1.1 Physcomitrella patens ....................................................................................... 23 1.3.1.2 Populus spp. ...................................................................................................... 24 1.4 Functional characterization of plant genes and gene products ..................................... 26 1.4.1 Bioinformatics........................................................................................................... 27 1.4.1.1 Phylogenetics .................................................................................................... 27 1.4.2 In vitro characterization ............................................................................................ 28 1.4.2.1 Heterologous production of proteins ................................................................ 29 1.4.2.2 Enzyme activity and substrate specificity of glycoside hydrolases .................. 29 1.4.2.3 Enzyme product analysis .................................................................................. 31 1.4.3 In planta and in situ characterization ........................................................................ 33 1.4.3.1 Transgenic techniques ....................................................................................... 33 1.4.3.2 Phenotype analysis ............................................................................................ 34 1.4.3.3 Gene expression analysis .................................................................................. 35 1.4.3.4 Protein localization ........................................................................................... 37 1.5 Research aims ............................................................................................................... 38 xi 1.6 Significance................................................................................................................... 40 Chapter 2: Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG16 and XTH proteins in plant lineages ..........................................................................................................................................41 2.1 Introduction ................................................................................................................... 41 2.2 Materials and methods .................................................................................................. 45 2.2.1 Data sources .............................................................................................................. 45 2.2.2 Identification of EG16s and XTHs ........................................................................... 46 2.2.3 Phylogenetic analysis ................................................................................................ 47 2.2.4 Expression data ......................................................................................................... 48 2.2.5 Accession numbers ................................................................................................... 48 2.3 Results and discussion .................................................................................................. 48 2.3.1 Cross-genomic census of EG16s and XTHs ............................................................. 48 2.3.2 EG16 and XTH homologs were not observed in the genomes of non‐plant eukaryotes ............................................................................................................................. 52 2.3.3 Red algal species have MLGase‐like sequences ....................................................... 54 2.3.4 A limited selection of green algae represents the most phylogenetically distant clades from angiosperms with canonical EG16 and XTH homologs ................................... 54 2.3.5 EG16 homologs were observed in many hornwort and liverwort transcriptomes.... 56 2.3.6 EG16 homologs are widespread in mosses, lycophytes, and ferns ........................... 57 2.3.7 EG16 homologs have been widely maintained among gymnosperms ..................... 58 2.3.8 EG16 homologs were variably maintained among angiosperms .............................. 61 xii 2.3.9 The molecular phylogeny of EG16 orthologs is generally concordant with contemporary estimates of Viridiplantae evolution .............................................................. 63 2.3.10 Observations on the putative biological roles of EG16 and EG16‐2 proteins ...... 67 2.4 Conclusions ................................................................................................................... 68 Chapter 3: Conservation of endo-glucanase 16 (EG16) activity across highly divergent plant lineages ................................................................................................................................69 3.1 Introduction ................................................................................................................... 69 3.2 Materials and methods .................................................................................................. 72 3.2.1 Materials ................................................................................................................... 72 3.2.2 Bioinformatics........................................................................................................... 72 3.2.3 Cloning of PpEG16 targets, transformation, protein production and purification ... 73 3.2.4 Enzyme kinetics ........................................................................................................ 76 3.2.5 Enzyme product analysis .......................................................................................... 77 3.2.6 Biomechanical assays ............................................................................................... 78 3.3 Results ........................................................................................................................... 78 3.3.1 PpEG16 is representative of non-vascular EG16s .................................................... 78 3.3.2 PpEG16 is predominantly a mixed-linkage -glucanase .......................................... 81 3.3.3 EG16 orthologs weaken plant cell walls ex vivo ...................................................... 85 3.4 Discussion ..................................................................................................................... 89 Chapter 4: Physcomitrella patens endo-glucanase 16 is involved in the cell wall development of young tissue ..............................................................................................................................93 4.1 Introduction ................................................................................................................... 93 4.2 Materials and methods .................................................................................................. 95 xiii 4.2.1 Data sources .............................................................................................................. 95 4.2.2 Growth conditions ..................................................................................................... 96 4.2.3 Plasmid construction and moss transformation ........................................................ 96 4.2.4 Genotyping ................................................................................................................ 97 4.2.5 Growth assays ........................................................................................................... 98 4.2.6 Sample preparation and microscopy procedures ...................................................... 98 4.3 Results ......................................................................................................................... 100 4.3.1 PpEG16 is expressed in young tissues.................................................................... 100 4.3.2 Growth phenotyping of ΔPpEG16 transgenics ....................................................... 100 4.3.3 PpEG16 localizes to the cell wall at the tips of protonema .................................... 105 4.4 Discussion ................................................................................................................... 109 Chapter 5: Functional analysis of Populus endo-glucanase 16 (EG16) ................................113 5.1 Introduction ................................................................................................................. 113 5.2 Materials and methods ................................................................................................ 115 5.2.1 Data sources ............................................................................................................ 115 5.2.2 Poplar growth conditions ........................................................................................ 115 5.2.3 gDNA extraction ..................................................................................................... 115 5.2.4 Cloning .................................................................................................................... 116 5.2.5 Plant transformation ................................................................................................ 117 5.2.6 Genotyping .............................................................................................................. 117 5.2.7 RNA extraction, cDNA preparation and RT-qPCR ................................................ 117 5.2.8 Tissue sectioning ..................................................................................................... 118 5.2.9 Promoter-GUS analysis .......................................................................................... 118 xiv 5.2.10 Antibodies ........................................................................................................... 118 5.2.11 Western blot ........................................................................................................ 119 5.2.12 Immunofluorescence ........................................................................................... 119 5.2.13 Microscopy ......................................................................................................... 120 5.2.14 Growth assay ....................................................................................................... 121 5.2.15 Soluble sugar analysis ......................................................................................... 121 5.2.16 MALDI-imaging ................................................................................................. 121 5.3 Results ......................................................................................................................... 123 5.3.1 PtEG16 expression is highest in young and expanding tissues .............................. 123 5.3.2 Populus EG16 immunolocalization ........................................................................ 126 5.3.3 Populus EG16 downregulation does not affect plant growth ................................. 127 5.3.4 Analysis of PtEG16 substrates in P39 tissues......................................................... 130 5.4 Discussion ................................................................................................................... 133 Chapter 6: General conclusions and future directions ...........................................................137 6.1 General conclusions .................................................................................................... 137 6.2 Future directions ......................................................................................................... 139 Bibliography ...............................................................................................................................142 Appendices ..................................................................................................................................158 Appendix A Supporting information for Chapter 2: Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG16 and XTH proteins in plant lineages ........................................................................................ 158 Appendix B Supporting information for Chapter 3: Conservation of endo-glucanase 16 (EG16) activity across highly divergent plant lineages .......................................................... 162 xv Appendix C Supporting information for Chapter 4: Physcomitrella patens endo-glucanase 16 is involved in the cell wall development of young tissue ....................................................... 172 Appendix D Supporting information for Chapter 5: Functional analysis of Populus endo-glucanase 16 (EG16) ............................................................................................................... 180 xvi List of Tables Table 3.1. PpEG16 and VvEG16 apparent Michaelis-Menten constants for polysaccharide hydrolysis ...................................................................................................................................... 84 Table B.1. Primer sequences used for ligation-independent cloning into pMCSG53 ................ 162 Table B.2. Primer sequences used for site-directed mutagenesis ............................................... 162 Table C.1. Primers used in this study ......................................................................................... 172 Table D.1. Primers used in this study ......................................................................................... 180 xvii List of Figures Figure 1.1. Molecular model of type I (A) and type II (B) primary cell walls found in angiosperms .................................................................................................................................... 2 Figure 1.2. Eukaryote phylogeny and occurrence of cell wall polysaccharides ............................. 4 Figure 1.3. Hemicellulose structures .............................................................................................. 8 Figure 1.4. Phylogeny of glycoside hydrolase family 16 ............................................................. 13 Figure 1.5. Enzymatic activities of xyloglucan endotransglycosylase/hydrolase (XTH) members....................................................................................................................................................... 15 Figure 1.6. Plant GH16 phylogeny ............................................................................................... 17 Figure 1.7. Non-classical secretory pathways ............................................................................... 18 Figure 1.8. Maximum likelihood phylogenetic tree of EG16 homologs ...................................... 19 Figure 1.9. The life cycle of the moss Physcomitrella patens ...................................................... 24 Figure 1.10. Populus alba x Populus grandidentata (P39) hybrid poplar plant ........................... 26 Figure 1.11. Michaelis-Menten model .......................................................................................... 31 Figure 1.12. MLG hydrolysis site of VvEG16 and other MLG endo-glucanases ......................... 33 Figure 2.1. Proposed evolution of glycoside hydrolase family 16 (GH 16) enzymes .................. 42 Figure 2.2. Comparison of bacterial mixed‐linkage endo‐glucanases, plant endo‐glucanases (EG16), and plant xyloglucan endotransglycosylase/hydrolases (XTHs) from glycoside hydrolase family 16 (GH16) ......................................................................................................... 43 Figure 2.3. Frequency of EG16 and XTH gene products among plants and marine eukaryotes.. 50 Figure 2.4. Aligned sequence logos of canonical EG16 and XTH sequences .............................. 51 Figure 2.5. Frequency of other GH16 gene products among plants and other eukaryotes ........... 53 xviii Figure 2.6. Quantitative analysis of EG16, XTH, and other GH16 homologs in translated genomes of plants and marine eukaryotes .................................................................................... 60 Figure 2.7. Maximum‐likelihood phylogeny of EG16 members .................................................. 65 Figure 3.1. EG16 orthologs among land plant clades ................................................................... 71 Figure 3.2. PpEG16 sequence and structure analysis ................................................................... 80 Figure 3.3. Michaelis-Menten kinetics of PpEG16.9 ................................................................... 83 Figure 3.4. Product analysis of PpEG16.9 .................................................................................... 85 Figure 3.5. Extension assays under constant load with sensitized hypocotyl walls ..................... 87 Figure 3.6. Stress-strain analysis of cucumber hypocotyls ........................................................... 88 Figure 4.1. Growth assay of wild-type, ΔPpEG16, and PpEG16 complementation moss lines 102 Figure 4.2. Dark growth assay of wild-type and ΔPpEG16 moss lines ...................................... 104 Figure 4.3. Assessment of senescence of ΔPpEG16 and wild-type plants ................................. 105 Figure 4.4. Confocal microscopy of PpEG16-mCherry localization in cell walls of protonema cell tips ........................................................................................................................................ 107 Figure 4.5. Confocal immunofluorescence microscopy of protonema treated with anti-xyloglucan antibodies (LM15) and anti-MLG antibodies (BG1) .................................................................. 108 Figure 5.1. Promoter-β-glucuronidase assays of transverse sections of P39 poplar lines expressing pPtEG16::GUS ......................................................................................................... 125 Figure 5.2. Confocal microscopy of immunolabelled P39 poplar (A) and EG16-downregulated (RC8-6) (B) sections ................................................................................................................... 128 Figure 5.3. Phenotyping of EG16-downregulated lines.............................................................. 130 Figure 5.4. Analysis of PtEG16 substrate xyloglucan in P39 tissues ......................................... 132 Figure A.1. Sequence logos of EG16‐2 motifs compared with EG16 motifs............................. 158 xix Figure A.2. Expression levels of PtEG16 in Populus trichocarpa tissues from the eFP browser..................................................................................................................................................... 159 Figure A.3. Expression levels of PaEG16 in Pinus abies tissues from ConGenie ..................... 160 Figure A.4. Expression of PpEG16 in Physcomitrella patens tissues from the eFP browser .... 161 Figure B.1. PpEG16 (Pp3c21_9420V3) mRNA transcripts aligned to chromosome 2 ............. 163 Figure B.2. PpEG16.9 homology model .................................................................................... 163 Figure B.3. Recombinant production of PpEG16 protein variants ............................................. 164 Figure B.4. Substrate screening of PpEG16 variants ................................................................. 165 Figure B.5. Temperature (A) and pH (B) profiles of PpEG16.9 using 1 mg·mL-1 MLG .......... 166 Figure B.6. PpEG16.9 stability at 4°C using different enzyme dilution buffers ........................ 167 Figure B.7. PpEG16.9 limit digest products ............................................................................... 168 Figure B.8. Activity of EG16 orthologs on AZCL-polysaccharides .......................................... 169 Figure B.9. Matching enzyme activities of E-XEGP and VvEG16 using AZCL-conjugated HEC and xyloglucan ............................................................................................................................ 170 Figure B.10. Stress-strain analysis for EG16 orthologs normalized to xyloglucanase activity of VvEG16 ....................................................................................................................................... 171 Figure B.11. Effect of EDTA and DTT on the stress-strain analysis of cucumber hypocotyl walls with PpEG16 ............................................................................................................................... 171 Figure C.1. EG16 expression in Physcomitrella patens tissues.................................................. 173 Figure C.2. Genotyping of Physcomitrella patens transgenic lines............................................ 174 Figure C.3. Representative growth progress of wild-type and ΔPpEG16 moss lines ................ 175 Figure C.4. Replicate growth assays of colonies of wild-type (wt) and two lines of PpEG16-deletion mutants .......................................................................................................................... 176 xx Figure C.5. Confocal live-imaging microscopy of PpEG16-mCherry expressing line and wild-type .............................................................................................................................................. 177 Figure C.6. Whole-mount immunostaining confocal microscopy of 7-day old wild-type and PpEG16-mCherry tissues............................................................................................................ 178 Figure C.7. Calcofluor-white staining of wild-type P. patens protonema tissue ........................ 179 Figure D.1. EG16 expression in Populus trichocarpa tissues .................................................... 182 Figure D.2. Genotyping of hybrid poplar P39 transformants ..................................................... 183 Figure D.3. Western blot for anti-PtEG16 antibodies ................................................................ 184 Figure D.4. Immunofluorescence microscopy using anti-PtEG16(full-length) antibody .......... 185 Figure D.5. Design of PtEG16 peptide used as the antigen to produce the anti-PtEG16(peptide) antibody....................................................................................................................................... 186 Figure D.6. RT-qPCR of wild-type and EG16 RNAi lines ........................................................ 187 Figure D.7. Assessment of MLG in P39 poplar tissues .............................................................. 188 Figure D.8. MALDI-imaging spectra of xyloglucan oligosaccharides in wild-type (A) and RNAi-EG16 P39 hybrid poplar (B) ....................................................................................................... 189 Figure D.9. Soluble sugar analysis of leaves from 3 months old greenhouse plants .................. 190 xxi List of Supplementary Material Supplementary Material Information. Supplementary material descriptions ................................... ......................................................................................... Supplementary Material Information.pdf Supplementary Material 2.1. Interactive table in Microsoft EXCEL format of all EG16, XTH and other GH16 members identified by BLAST ........................................................... Suppl_2-1.xlsx Supplementary Material 2.2. MUSCLE alignment of EG16‐2 sequences with EG16s, XTHs and MLGases in FASTA format.................................................................................... Suppl_2-2.fasta Supplementary Material 2.3. MUSCLE alignment of β‐bulge‐containing GH16 sequences in FASTA format ........................................................................................................ Suppl_2-3.fasta Supplementary Material 2.4. MUSCLE alignment of red algae GH16 members in FASTA format................................................................................................................................. Suppl_2-4.fasta Supplementary Material 2.5 MUSCLE alignment of hornwort and basal eudicot EG16 members missing key catalytic residues in FASTA format ................................................... Suppl_2-5.fasta Supplementary Material 2.6. MUSCLE alignment of Ginkgo sequences with other EG16s, XTHs and MLGases in FASTA format ............................................................................. Suppl_2-6.fasta Supplementary Material 2.7. RAxML phylogeny in NEXUS format ....................... Suppl_2-7.nex Supplementary Material 2.8. Manually curated alignment in FASTA format used for maximum‐likelihood phylogeny of EG16 members ................................................................ Suppl_2-8.fasta Supplementary Material 2.9. Alignment of EG16 sequences in FASTA format used to produce sequence logos ........................................................................................................ Suppl_2-9.fasta Supplementary Material 2.10. Alignment of XTH sequences in FASTA format used to produce sequence logos ...................................................................................................... Suppl_2-10.fasta Supplementary Material 3.1. PpEG16 coding sequence MUSCLE alignment ...... Suppl_3-1.fasta xxii Supplementary Material 3.2. Homology model of PpEG16.9 ................................ Suppl_3-2.pdb Supplementary Material 3.3. MUSCLE alignment of E. coli-optimized coding sequences of PpEG16.1, PpEG16.9, and PpEG16.10 ................................................................. Suppl_3-3.fasta Supplementary Material 3.4. MUSCLE alignment of EG16 protein sequences from PtEG16, VvEG16, and PpEG16 ............................................................................................ Suppl_3-4.fasta xxiii List of Abbreviations 1KP 1000 Plants database AA Auxiliary activity AGP Arabinogalactan protein ANA Plant group comprising members from orders Amborellales, Nymphaeales and Austrobaileyales ANOVA Analysis of variance AZCL Azo-dyed and azurine cross-linked BI Bayesian inference BCA Bicinchoninic acid BLAST Basic local alignment search tool bp Nucleic acid base pairs BSA Bovine serum albumin CAZy Carbohydrate active enzyme database CAZymes Carbohydrate-active enzymes CBM Carbohydrate-binding modules cDNA Complementary deoxyribonucleic acid CE Carbohydrate esterase CES Cellulose synthase CFP Cyan fluorescent protein cGM Carob galactomannan CMC Carboxymethyl cellulose CSL Cellulose synthase-like xxiv Da Dalton ddH2O Sterile ultra-pure water DNA Deoxyribonucleic acid DTT Dithiothreitol EC Enzyme Commission EDTA Ethylenediaminetetraacetic acid eFP Electronic fluorescent pictograph browser EG Endo-glucanase G β(1,4)-linked glucose G3 β(1,3)-linked glucose Gal Galactose GAX Glucuronoarabinoxylan gDNA Genomic deoxyribonucleic acid GH Glycoside hydrolase GH## Glycoside hydrolase family ## Glc Glucose GlcA Glucuronic acid GT Glycosyltransferase GUS β-glucuronidase HEC Hydroxyethyl cellulose HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HG Homogalacturonan xxv HPAEC-PAD High-performance anion-exchange chromatography with pulsed amperometric detection HPLC High-performance liquid chromatography ID Sequence identity ISH in situ hybridization ITO Indium-tin-oxide IPTG Isopropyl β-D-1-thiogalactopyranoside JGI Joint Genome Institute JTT Jones-Taylor-Thornton algorithm kDa Kilodalton kGM Konjac glucomannan L β(1,2)-galactosyl-α(1,6)-xylosyl-β(1,4)-glucose LB Lysogeny broth LIC Ligation independent cloning m/z Mass-to-charge ratio M β(1,4)-linked mannose MALDI-TOF Matrix-assisted laser desorption ionization coupled to time-of-flight mass spectrometer Man Mannose ML Maximum‐likelihood phylogeny MLG Mixed-linkage β(1,3)/β(1,4)-glucan MLGases Bacterial β(1,3)/β(1,4) mixed‐linkage endo‐glucanases MLGOs Mixed-linkage glucan-derived oligosaccharides xxvi MOPS 3-(N-morpholino)propanesulfonic acid mRNA Messenger ribonucleic acid MUSCLE Multiple sequence comparison by log-expectation MS Mass spectrometry mya Million years ago NA Numerical aperture NAA 1-naphthaleneacetic acid NCBI National Center for Biotechnology Information ND Not determined OD Optical density NJ Neighbor joining PCR Polymerase chain reaction PDB ID Protein Data Bank identifier PDB Protein Data Bank PEG Polyethylene glycol PL Polysaccharide lyase PLL Poly-L-lysine RG-I Rhamnogalacturonan-I RG-II Rhamnogalacturonan-II RMA Robust multi-array average RNAi Ribonucleic acid interference RPM Revolutions per minute RT-qPCR Quantitative reverse transcription-polymerase chain reaction xxvii SAR Plant taxonomic supergroup containing members of stramenopiles (heterokonts), alveolates and Rhizaria SDS-PAGE Sodium dodecyl sulphate-polyacrylamide gel electrophoresis SEC Size-exclusion chromatography TBST Tris-buffered saline with Tween-20 detergent TD Transmitted light differential interference contrast images TE Tris-buffered saline with ethylenediaminetetraacetic acid TEV Tobacco etch virus TRIS Tris(hydroxymethyl)aminomethane wAX Wheat arabinoxylan WPM Woody plant media wt Wild-type X α(1,6)-xylosyl-β(1,4)-glucose XEH Xyloglucan endohydrolase XET Xyloglucan endotransglycosylase XTH Xyloglucan endotransglycosylase/hydrolase XyG Xyloglucan XGO Xyloglucan-derived oligosaccharide Xyl Xylose XXT Xyloglucan xylosyl transferases xxviii Acknowledgements I would like to give my gratitude to my Ph.D. supervisor, Professor Harry Brumer. I thank him for giving me the opportunity to be involved in various fields, for helping me improve my writing and presentation skills, and for supporting my progress and independence. I would also like to thank Professor A. Lacey Samuels for giving me vital support in my experiments, including me as part of her lab, and for her continual professional support. My inclusion in the Samuels lab exposed me to unique opportunities for learning and development. I greatly thank Professor Shawn D. Mansfield for allowing me to work in his lab, participate in his lab meetings, and for his support of my research. I thank my supervisory committee member, Professor Nobuhiko Tokuriki, for his direction and advice. I thank the funding sources, NSERC-CREATE/DFG-IRTG program Plant Responses To Eliminate Critical Threats (PRoTECT), Michael Smith Laboratories staff, and UBC Biochemistry and Molecular Biology department and staff. In particular, I would like to thank Doris Metcalf for her support during my degree. I am appreciative of UBC Bioimaging facility staff Dr. Miki Fujita, Kevin Hodgson, and Dr. Eunkyoung Lee for assistance with microscopy experiments and UBC Greenhouse staff, Melina Biron and Taylor Scriber, for their help with poplar growth trials. I thank Professor Ljerka Kunst and Dr. Tegan Haslam (UBC Botany) for providing Physcomitrella plants and related resources, and Professor Magdalena Bezanilla (Dartmouth College, Biological Sciences) and Dr. Gillian Dean (UBC Botany) for their helpful suggestions. I want to acknowledge the Brumer lab members, past and present. I am grateful to Dr. Shaheen Shojania for his general support and for providing tasty vegan birthday treats. I express my gratitude to James Li for his help with HPLC experiments and his support and Dr. Kazune Tamura for valuable discussions and assistance with enzymology. I thank Dr. Nicholas McGregor xxix for his assistance with protein work and bioinformatic analyses, Dr. Alexander H. Holm for computational assistance, Dr. Changqing Wang for his help with MALDI-TOF, and Drs. Julie Grondin, Jonathan Briggs, Gregory Arnal, and Laleh Solhi for helpful discussions. Special thanks to Sean McDonald for his continual support and help with figure making, enzymology experiments, and writing. I am tremendously grateful to Dr. Namrata Jain for invaluable conversations, friendship, and encouragement. I offer my gratitude to all Samuels lab members, past and present, especially Dr. Miranda Meents and Dr. Sam Livingston, for their guidance with antibody work and microscopy, and Dr. Mathias Schuetz, Mendel Perkins, and Natalie Hoffmann for helpful discussions. Special thanks to Kenzie Arnott and Jan Xue for their continual support and friendship. I would also like to extend my gratitude to the Mansfield lab members for their hospitality. I thank Dr. Faride Unda and Yaseen Mottiar for their support with poplar work and HPLC, Michelle von Loessl for her assistance with promoter-GUS analyses, Dr. Eliana Gonzales-Vigil for her help with RT-qPCR, and Michael Bilek for assistance with HPLC. I appreciate Dr. Letitia Da Ros, Xinyi Huang, Elizabeth Mahon, and Pablo Antonio Chung for helpful discussions. I would like to express profound gratitude to my parents, Tami and Haim Behar, for their boundless love, encouragement, and support. I am immensely appreciative of my sisters, Gali and Tal Behar, for their motivation and reassurance. I thank my grandparents, Larissa Bagdasarian, extended family, and my sisters’ families for their backing. I thank my cats Panda, June, and Rosie for their company and support. I am grateful to my friends for their support (who have not been mentioned above already), Eitan, Topher, Josh, Gina, Sina, Dana, Maria, Anne, Ravi, and Joey. Finally, I give my highest gratitude to Alex Migolin for his unfaltering encouragement, support, and advice during my degree. I couldn’t have done it without you. xxx Dedication To my family. Tami, Haim, Gali, Tal, and Alex. 1 Chapter 1: Introduction The dynamic modification of the plant cell wall polysaccharides is carried out by carbohydrate active-enzymes such as xyloglucan endotransglycosylase/hydrolase (XTH). The plant endo-glucanase 16 (EG16) enzymes were recently identified as a unique clade that is structurally and functionally intermediate between the bacterial mixed-linkage β(1,3)/(1,4)-glucanases and the plant XTHs in glycoside hydrolase family 16 (GH16). Although EG16 gene product activity and structure have been demonstrated, their biological role is unknown. This thesis employs a multi-disciplinary approach to the characterization of the newly designated EG16 group and their biological functions in plant cell walls. 1.1 Plant cell walls Plantae represents one of the major taxonomic kingdoms whose members are essential to the environment as primary energy producers. Plants are photosynthetic eukaryotes that grow continuously throughout their lifespans. To accomplish this, plants rely on a complex structure surrounding their cells known as the cell wall. Evidence suggests that the cell wall has a myriad of functions, including those related to the regulation of growth and shape, protection from mechanical and osmotic stress, and defence and detection of pathogens (Evert and Esau, 2006). In some plant cells, further cell wall specialization allows for additional functions such as water transport, enhanced structural support, and seed protection. Cell walls are dynamic and change throughout the plant lifecycle and often in response to the changes in the environment (Carpita et al., 2001, Jamet and Dunand, 2020). The cell wall comprises two distinct layers known as primary and secondary cell walls. The primary cell wall forms in the cell plate as part of cell division. These can grow and expand up to 1000-fold (Buchanan et al., 2000). Following primary cell wall production, secondary cell walls form in 2 specialized cells (Buchanan et al., 2000). Polysaccharide components, including cellulose, pectin and hemicelluloses, comprise the majority of the plant primary cell wall content (Figure 1.1). Figure 1.1. Molecular model of type I (A) and type II (B) primary cell walls found in angiosperms (reused with permission from (Buchanan et al., 2000)). Both cell wall types contain cellulose microfibrils coated by and crosslinked by hemicelluloses embedded in a pectin matrix. (A). Type I cell walls contain xyloglucan strands and small amounts of other hemicelluloses in a homogalacturonan and rhamnogalacturonan I matrix. (B). Type II cell walls are exclusive to commelinid monocots and contain cellulose, glucuronoarabinoxylan as the dominant crosslinking glycan, and pectins. In cereals (Poales), mixed linkage β-glucan is also present in more significant amounts as the plant is growing. 1.1.1 Plant cell wall types The primary cell walls of the taxonomically grouped plant species, bryophytes, pteridophytes, gymnosperms, and angiosperms, are generally described by polysaccharide content, while some are also assigned a cell wall type designation. However, researchers have expressed skepticism about the utility and accuracy of this categorization (Leroux et al., 2015). Indeed, cell 3 wall types are highly simplified portrayals of the primary cell walls, as they differ between cells, tissues, stages of development, and exposure to various environmental circumstances (Popper et al., 2011, Leroux et al., 2015). Still, as general descriptors, primary cell wall categories assist in discussing and understanding the wall contents of distinct plant groups. In this section, broad plant cell wall designations will be described, followed by detailed explanations of the molecular structure of cell wall components in the subsequent sections. The primary cell wall architecture of angiosperms is divided into Type I and Type II (Figure 1.1). Type I cell walls are present in most angiosperms, including members of Brassicaceae, e.g., thale cress (Arabidopsis thaliana) and Solanaceae, e.g., tomato (Solanum lycopersicum), but they also generally describe gymnosperm primary cell walls (O'Neill and York, 2003). Type I primary cell walls are described as a network of crystalline cellulose microfibrils embedded in a matrix of amorphous polysaccharides, consisting mainly of pectins along with hemicelluloses, such as xyloglucan (Figure 1.1A). Furthermore, some eudicot cell walls contain large amounts of proteins, which interact with pectins or interlink with each other to form networks (Buchanan et al., 2000). Type II cell walls describe the walls of commelinid monocots, e.g., date palm (Areca catechu), which are comprised of cellulose microfibrils, glucuronoarabinoxylan, and low amounts of both pectin and structural proteins (Carpita and Gibeaut, 1993) (Figure 1.1B). Members of the Poales, an order within the commelinid monocots, also contain mixed-linkage β(1,3)/β(1,4)-glucan (MLG), that coat the cellulose microfibrils in tissues of growing plants (Figure 1.1B). Primary cell walls of non-angiosperm plant groups vary but retain the main framework of cellulose microfibrils and amorphous polysaccharides (Popper et al., 2011). The cell walls of charophyte green algae, bryophytes and ferns, contain the matrix polysaccharide xyloglucan, 4 mannan, xylan, as well as the pectic polysaccharides homogalacturonan (HG) and rhamnogalacturonan 1 (RG-1) (Popper et al., 2011, Domozych et al., 2012, Roberts et al., 2012). Figure 1.2. Eukaryote phylogeny and occurrence of cell wall polysaccharides (reused with permission from Popper, Z.A., et al. 2011). Cellulose synthase (Ces) and cellulose synthase-like (Csl) gene families are noted on branches. PE – primary endosymbiosis, SE – secondary endosymbiosis. 5 MLG is also found in chlorophyte green algae and some pteridophyte species, notably those of the horsetail family (Equisetaceae) (Xue and Fry, 2012, Leroux et al., 2015) (Figure 1.2). Ferns, another pteridophyte group, have Type III cell walls, which are cell walls with a higher amount of mannan and a lower amount of pectin (Silva et al., 2011). Glucomannan makes up the majority of the hemicelluloses in gymnosperms, containing up to 10% of their total cell wall (w/w) (Popper and Fry, 2003). 1.1.2 Plant cell wall polysaccharides As discussed above, polysaccharides play a central role in plant physiology. The primary cell wall contains several classes of polysaccharides and other components. Their molecular structure and physiology are discussed in the following subsections. 1.1.2.1 Cellulose Cellulose is a ubiquitous plant polysaccharide that forms the majority of the plant cell wall. In fact, cellulose makes up 20-30% of primary cell walls and roughly 50% (or more in specialized cell walls) in secondary cell walls (Buchanan et al., 2000, O'Neill and York, 2003). Cellulose has a linear structure of long chains of β(1,4)-linked glucosyl residues that adhere to other chains via hydrogen bonds to form paracrystalline microfibrils of 18 strands (McNamara et al., 2015, Kubicki et al., 2018). Cellulose microfibrils provide high tensile strength and serve as the load-bearing polysaccharide (McFarlane et al., 2014). Moreover, cellulose microfibrils regulate cell growth, where their orientation is transverse to the direction of growth (Thomas et al., 2013). 1.1.2.2 Pectin Pectins are a category of complex cell wall polysaccharides that include homogalacturonan (HG), rhamnogalacturonan-I (RG-I), and rhamnogalacturonan-II (RG-II) (Saffer, 2018). HG and 6 RG-II have an α(1,4)-linked galacturonic acid backbone. HG is linear with methyl esterification, while RG-II is highly branched with a non-repeating pattern of arabinose, rhamnose, galactose, fucose, and other saccharide units. RG-I has a backbone of alternating galacturonic acid and rhamnose residues, has branches of arabinans and galactans, and can sometimes be substituted with fucose (Saffer, 2018). Pectins provide a gelatinous matrix for the primary cell wall architecture and are involved in cell expansion and cell adhesion (Palacio-Lopez et al., 2020). Experimental evidence has demonstrated an essential role for pectins. Mutations to HG synthesis machinery in A. thaliana cause a dwarf phenotype (Saffer, 2018). In addition, plants with fucose-deficient mutations affecting RG-II structure have delicate tissues, and mutations affecting rhamnose synthesis that lead to reduced amounts of RG-I have cell expansion defects (Saffer, 2018). Moreover, research suggests that pectins have some functional redundancy with hemicelluloses, xyloglucan in particular, and interacts with cellulose surfaces (Park and Cosgrove, 2015). 1.1.2.3 Hemicellulose Hemicelluloses are heterogeneous polysaccharides present in the cell walls of plants that share a β(1,4)-linked sugar backbone with an equatorial configuration at C1 and C4. Hemicelluloses were initially defined as the remaining polysaccharides after the extraction of pectin, which can be isolated with an alkaline reagent application (Bauer et al., 1973, Ghosh et al., 1985). The hemicellulose class of polysaccharides includes xyloglucan, xylans, mannans, and MLG (Scheller and Ulvskov, 2010) (Figure 1.3). These are present in all land plants except for MLG, which has been observed only in the Poales and Equisetum species (Popper et al., 2011). Several models aim to explain the function of hemicelluloses. The multicoat model describes the formation of hemicellulose layers around cellulose microfibrils (Scheller and 7 Ulvskov, 2010). The sticky network model illustrates hemicellulose as crosslinkers or tethers between cellulose microfibrils via non-covalent bonds. The stratified wall model was also considered, which is a fusion of the two models (Cosgrove, 2000). However, many researchers note that these simplified representations are not adequate for explaining cell wall architecture and the function of hemicellulose in particular (Cosgrove, 2018). Emerging roles have been proposed, particularly for xyloglucan, the more prominent hemicellulose in angiosperm walls, which will be described in its own section. Uncontroversially, hemicelluloses act as storage molecules serving as a readily available energy source for germinating seeds (Scheller and Ulvskov, 2010). In human society, hemicelluloses are essential food sources found in high amounts in many crops. For example, MLG is abundant in many cereal grains and consumption is positively associated with better gut health (Tamura et al., 2017). 1.1.2.3.1 Mannans Mannan and galactomannans have a β(1,4)-linked mannosyl backbone, while glucomannans contain additional β(1,4)-linked glucosyl residues distributed in the backbone as well (Scheller and Ulvskov, 2010) (Figure 1.3). Mannans are most abundant in non-angiosperm plant cell walls, particularly in charophyte green algae, mosses, lycophytes and ferns, as well as in the secondary cell walls of gymnosperms. Mannans have a vital role in embryo development and as seed storage polysaccharides (Scheller and Ulvskov, 2010). Mannans also play a role as prominent cell wall polysaccharides in some plant species, as described above. The differing observations of mannan suggest various biological functions depending on the phylogeny of the organism and the tissue type (Silva et al., 2011). 8 Figure 1.3. Hemicellulose structures (reused with permission from Scheller and Ulvskov, 2010). 9 1.1.2.3.2 Xylans Xylans are polysaccharides that have a β(1,4)-linked xylosyl backbone (Figure 1.3). Xylans are categorized into three groups according to their branching patterns. Glucuronoxylans have branches of α(1,2)-linked glucuronosyl residues common in the secondary cell walls of eudicots (Scheller and Ulvskov, 2010). Arabinoxylans and glucuronoarabinoxylans are substituted with arabinose residues and are common in commelinid monocots. Gymnosperms cell walls are also rich in arabinoxylans (Scheller and Ulvskov, 2010). Xylans contribute to the growth and development of plants by providing rigidity and protection from pathogens (Buchanan et al., 2000). Additionally, xylans form an important class of polysaccharides that are of value to humans. Xylans are found in high amounts in cereal grains, and they are also used in various high-value industries. For example, the xylose component of xylans is utilized for xylitol production. A product that is used as a sugar substitute, and as a cavity-reducing agent (Saha and Bothast, 1997). 1.1.2.3.3 Xyloglucan Xyloglucans were initially discovered in the late 1950s in nasturtium (Tropaeolum majus), a flowering plant from the Tropaeolaceae family and other plant seeds (Stratilová et al., 2020b). Since then, the universal nature of xyloglucan in plant families has been illustrated by the presence of associated genetic machinery in green algae and all land plants, such as cellulose synthase-like family C (CSLC) and xyloglucan xylosyl transferases (XXT) genes (Del-Bem, 2018). Xyloglucan is a polysaccharide composed of repeating units of β(1,4)-linked glucosyl residues with short side chains of xylosyl, galactosyl, fucosyl, and arabinosyl residues (Schultink et al., 2014) (Figure 1.3). It was shown to have a wide array of diverse branching patterns dependent on plant species (Schultink et al., 2014). Xyloglucan molecules follow a repeating motif of xylosylation, typically XXXGn or XXGGn, with varying sidechains on the xylosyl residues. 10 The most widely accepted paradigm of the primary cell walls describes a role for xyloglucan as a load-bearing polysaccharide, which coats and crosslinks between cellulose microfibrils (Figure 1.1). However, there have been questions about the biological function of xyloglucan, as several cases put doubt into the current model. Some plants, such as celery (Apium graveolens), have small amounts of xyloglucan or any other hemicellulose (Scheller and Ulvskov, 2010). In addition, knocking out the genes required for xylosylation in A. thaliana effectively eliminates the ability of the plant to synthesize xyloglucan, resulting in small plants that grow normally otherwise (Cavalier et al., 2008, Kim et al., 2020). However, the knockout of certain xyloglucan side chains leads to significant growth phenotypes, such as the dwarf mutant in A. thaliana when galactosylation is inhibited (Kong et al., 2015, Park and Cosgrove, 2015). Additionally, overexpression of a xyloglucanase causes plants to grow larger, supporting the theory that xyloglucan has a role in growth regulation (Park et al., 2004). The examples above show that the role of xyloglucan as a load-bearing polysaccharide does not adequately explain its entire function. Recent proposals have outlined a new role, specifically for xyloglucan. The hemicellulose is proposed to be a “spacer” to separate cellulose microfibrils, as the polysaccharide is not fully expanded in the walls (Cosgrove, 2018, Kim et al., 2020). In addition, it has been suggested that xyloglucan forms “biomechanical hot spots” that bind two adjacent cellulose microfibrils to create load-bearing junctions (Park and Cosgrove, 2012). Lastly, xyloglucan may serve different roles depending on the plant species and tissues. 1.1.2.3.4 Mixed-linkage β-glucan MLG is a linear, kinked polysaccharide made up of β(1,4) and β(1,3)-linked glucosyl residues (Figure 1.3) that acts as a hemicellulose in the cell walls of some plants and other organisms, as 11 well as a seed storage carbohydrate (Lim et al., 2020). MLG was identified in various organisms, including fungi, brown algae, red algae, green algae, Equisetum spp. and most notably in gramineous monocots (Popper et al., 2011) (Figure 1.2). The presence of MLG in disparate and distantly related organisms can potentially be explained by convergent evolution (Popper and Tuohy, 2010). It is a possibility that MLG plays various roles due to the difference in the way it occurs in the plant. In the Poales, MLG is abundant in growing tissues and decreases later in development, suggesting a role in expansion and growth (Kiemle et al., 2014). In Equisetum spp., a wall strengthening role has been suggested, as the polysaccharide concentration is higher in mature tissues of the plant (Fry et al., 2008). Although the precise function of MLG is not known, its role has been examined by genetic manipulation. Knockout of CslF6, the glycosyltransferase implicated in MLG synthesis, in rice (Oryza sativa), causes the plants to be significantly shorter but otherwise grow normally (Vega-Sánchez et al., 2012). Overexpression of CslF6, implicated in MLG synthesis in barley (Hordeum vulgare L.), negatively affects grain development (Lim et al., 2020). As with manipulations of xyloglucan, it appears that plants have a robust ability to persevere with significant changes to their cell walls. 1.1.3 Cell wall proteins Cell wall proteins are implicated in structural architecture, maintenance, and modification of the wall as development progresses. In Arabidopsis, as many as 400 different proteins have been observed in the cell wall (Carpita et al., 2001). Structural proteins such as extensins and arabinogalactan proteins (AGPs) are implicated in plant development, elongation, and signalling in the latter group (Jamet and Dunand, 2020). For example, expansins are a group of nonenzymatic proteins implicated in cell expansion, fruit ripening, and pollination (Cosgrove, 2016). 12 1.2 Carbohydrate-active enzymes (CAZymes) CAZymes are catalytic enzymes that degrade, modify or form glycosidic bonds. They are present in all domains of life (Pinard et al., 2015, Viborg et al., 2019). CAZymes are found in large numbers in plants and constitute 3-10% of their genes, more than any other taxa (Coutinho et al., 2003, Pinard et al., 2015). The complexity of the cell wall, as well as early genome duplications and local duplications of these genes, can explain this high proportion. 1.2.1 Carbohydrate-Active enZymes (CAZy) database The CAZy database (http://www.cazy.org) includes members from five enzyme classes: Glycoside Hydrolases (GHs), Glycosyltransferases (GTs), Polysaccharide Lyases (PLs), Carbohydrate Esterases (CEs) and Auxiliary Activities (AAs) (Cantarel et al., 2009). Carbohydrate-binding modules (CBMs), although non-enzymatic, are proteins that specifically bind carbohydrates and therefore are also included in the CAZy database (Lombard et al., 2014). Each class of enzyme and binding protein contains multiple subfamilies, the most prominent one being the GHs, comprising 171 subfamilies as of June 2021. The following sections will focus on GH16, as the family comprises the subjects of this thesis, the EG16 enzymes. 1.2.2 Glycoside hydrolase family 16 (GH16) GH16 includes members that are active on β(1,4) or β(1,3) linkages of glucans and galactans. GH16 enzymes are mainly found in bacterial and fungal species, and they share a common active site motif (subgroup 1: EXDXXE, subgroup 2: EXDXE), similar primary sequence, and tertiary structure (β-jelly roll fold) (Keitel et al., 1993, Planas, 2000) (Figure 1.4). GH16 enzymes are sister to GH7 endo-glucanases in clan GH-B that share similar structural features and catalytic machinery, but are phylogenetically distinct (in the CAZy classification, a “Clan” is a higher-order grouping of families, roughly equivalent to a “superfamily”) (Michel et 13 al., 2001). Plant GH16 comprises two groups endo-glucanase 16 (EG16) and xyloglucan endotranglycosylases/hydrolases (XTH) (Eklöf et al., 2013, McGregor et al., 2017). 1.2.2.1 Bacterial mixed-linkage glucanases (MLGases) Bacterial MLGases (also known as licheninases; EC 3.2.1.73) are GH16 enzymes present in some bacterial species (e.g., Bacillus spp.) that degrade β(1,4) linkages of MLG (Planas, 2000, Gaiser et al., 2006). MLGases form a sister group to the plant GH16 clade (Figure 1.4). Until the discovery of the EG16 enzymes in plants, which will be described below, bacterial MLGases were considered the closest evolutionary relatives to XTHs. Figure 1.4. Phylogeny of glycoside hydrolase family 16 (reused from Behar et al. 2018). GH16 is a polyspecific group of enzymes that share a similar sequence and protein structure. Most GH16 members are bacterial and fungal except for EG16 and XTH enzymes, which are plant members. 14 1.2.2.2 Xyloglucan endotransglucosylase/hydrolase (XTH) enzymes XTH enzymes are plant GH16 members that cleave or rearrange xyloglucan in the plant cell walls. The XTH family comprises two different types of enzymes, xyloglucan endotransglycosylases (XETs, EC 2.4.1.207) and xyloglucan endohydrolases (XEHs, EC 3.2.1.151). XETs rearrange xyloglucan by non-hydrolytic cleavage and ligation to other strands, and XEHs shorten xyloglucan chains via hydrolytic cleavage (Eklöf and Brumer, 2010, Viborg et al., 2019) (Figure 1.5). The XTH family members have been highly studied, from enzymatic research and structure determination to in vivo characterizations (Eklöf and Brumer, 2010, Yokoyama et al., 2010, Kaewthai et al., 2013). XTH genes have been observed in all land plant clades and are present in large groups of 20-60 homologs. For example, the genomes of A. thaliana, Populus trichocarpa and the moss Physcomitrella patens encode 33, 46, and 32 XTH orthologs, respectively (Rose et al., 2002, Eklöf and Brumer, 2010, Yokoyama et al., 2010). Phylogenetic analyses of XTH genes show a separation into three groups I/II, III-A, and III-B. Groups I/II and III-B comprise members that exhibit XET activity only, while group III-A members exhibit XEH activity (Eklöf and Brumer, 2010). XTH gene products have been implicated in cell wall loosening, cell wall strengthening, and integrating new xyloglucan polymers into the cell wall (Eklöf and Brumer, 2010). Numerous studies have shown that XTH gene products are generally involved in plant growth (Rose et al., 2002). However, some studies suggest that the biological roles of XTHs are diverse (Rose et al., 2002, Hayashi and Kaida, 2011, Cosgrove, 2016, Stratilová et al., 2020a). For instance, XTH22 expression in A. thaliana is associated with environmental stressors such as wind and wounding (Rose et al., 2002). In addition, many studies have shown that XTH activity is associated with fruit 15 ripening processes, which involve cell wall degradation (Rose and Bennett, 1999, Rose et al., 2002, Miedes and Lorences, 2009). Figure 1.5. Enzymatic activities of xyloglucan endotransglycosylase/hydrolase (XTH) members (adapted from Cosgrove, 2005 with permission). (A). Xyloglucan endotranglycosylases (XETs) cleave the backbone of xyloglucan strands and form a new glycosidic bond with another xyloglucan strand. (B). Xyloglucan endohydrolase (XEH) activity involves the hydrolysis of a xyloglucan backbone. 1.2.2.3 Endo-glucanase 16 (EG16) enzymes In 2013, several plant genes, designated as endo-glucanase 16 (EG16) enzymes, were first shown to form a group distinct from XTHs phylogenetically, structurally, and functionally (Eklöf et al., 2013). Prior to this finding, XTHs were the only known GH16 members in plants, with 16 bacterial MLGases being their closest relatives according to phylogenetic models (Eklöf and Brumer, 2010) (Figure 1.4, Figure 1.6). EG16 gene products, previously considered to be XTH, have a protein scaffold that is intermediate between bacterial MLGases and the XTHs (Eklöf et al., 2013) (Figure 1.6). The EG16 protein structure lacks both the conserved N-terminal loop extension found in bacterial MLGases, and the C-terminal extension of the XTH proteins (“XET_C” PFAM domain PF06955) (Figure 1.6). In addition to sequence and structural differences, the EG16 clade differs from the XTH family in the number of orthologs present per genome. Whereas EG16 genes are present once per plant genome, if at all, XTH orthologs are observed in a large number of copies in all plant genomes (Eklöf et al., 2013). Lastly, a significant difference between EG16 and XTH gene products is their substrate specificity. EG16 enzymes were found to hydrolyze MLG prominently, as well as xyloglucan and cellulosic substrates, whereas XTHs have a high specificity for xyloglucan, as described above (Eklöf et al., 2013, McGregor et al., 2017). Unlike XTH genes, EG16 genes do not encode a signal peptide, are not predicted to have N-glycosylated sites, and have several unbridged cysteine residues (Eklöf et al., 2013, McGregor et al., 2017). These features would suggest cytosolic localization of EG16, but this raises the question of how EG16 gene products would function without access to their demonstrated cell wall substrates. However, the possibility remains that EG16 enzymes may be transported via unconventional secretion mechanisms, which have been observed in mammalian, yeast, and plant cells (Agrawal et al., 2010, Ding et al., 2014) (Figure 1.7). 17 Figure 1.6. Plant GH16 phylogeny (adapted from Eklöf et al., 2013). EG16 orthologs have structural and functional characteristics that suggest that they are evolutionary intermediates between the bacterial MLGases and XTHs. The substrates of EG16 gene products from Populus trichocarpa and Vitis vinifera were MLG and xyloglucan. A census of EG16 orthologs showed that they were found in various plants, including the moss P. patens, lycophyte Selaginella moellendorffii, and several graminaceous monocots and eudicots for a total of 33 homologs out of ca. 40 genomes (McGregor et al., 2017) (Figure 1.8). Interestingly, the census revealed the absence of EG16 orthologs from many genomes, particularly those of eudicots, notably Arabidopsis spp. and Solanaceae members such as Solanum lycopersicum and Nicotiana spp. (Eklöf et al., 2013, McGregor et al., 2017) (Figure 1.8). Due to a lack of non-vascular genomes and transcriptomes, the distribution of EG16 genes across plant lineages was not fully resolved (Eklöf et al., 2013, McGregor et al., 2017). 18 Figure 1.7. Non-classical secretory pathways (reused from (Agrawal et al., 2010)). Cytosolic protein secretion mechanisms observed in mammalian cells. Activity assays of EG16 enzymes from black cottonwood (P. trichocarpa; PtEG16) and grapevine (Vitis vinifera; VvEG16) show that they employ a unique cleavage pattern in MLG digestion distinct from characterized MLGases and plant GH17 MLG endohydrolases (Fincher, 2009, McGregor et al., 2017). Characterized EG16 gene products were demonstrated to cleave the β(1,4) linkage before the β(1,3)-linked glucose (non-reducing side) producing glucose, cellobiose, and β-D-Glcp-(1,3)-β-D-Glcp-(1,4)-β-D- Glcp-(1,4)-β-D-Glcp (G3GGG) in an MLG limit digest (Eklöf et al., 2013, McGregor et al., 2017). Two studies have examined the factors that influence EG16 expression. The EG16 homolog in P. patens (PpEG16; previously XTH32) was highly expressed compared to XTH16 and XTH21 that were tested alongside (Yokoyama et al., 2010). PpEG16 expression was observed in the young protonema tissue, and young stems and phyllids (leaf-like structures) of gametophores (Yokoyama et al., 2010). Regulation of EG16 from P. trichocarpa (PtEG16; previously XTH8) 19 was shown to be slightly downregulated by several hormonal treatments, including 6-benzylaminopurine, indole-3-acetic acid, salicylic acid, and brassinosteroids (Ye et al., 2012). Conversely, PtEG16 was slightly upregulated by jasmonic acid and abscisic acid (Ye et al., 2012). Figure 1.8. Maximum likelihood phylogenetic tree of EG16 homologs (adapted from (McGregor et al., 2017)). 33 EG16 protein sequences were identified from GenBank and Phytozome and aligned with XTH sequences as outliers and MLGase (licheninase) sequences as a root. 20 The studies discussed in this section designated EG16 as a distinct group, with protein structure and enzymatic function separate from XTH gene products. However, the EG16 family is still poorly understood. In particular, the EG16 gene distribution across different plant families is not known. Additionally, it is not clear whether non-angiosperm homologs exhibit similar enzyme activity, where the EG16 gene product localizes, and its biological function. Sections 1.3 and 1.4 will outline the model organisms used in this work, and review the approaches and techniques utilized in the characterization of the EG16 members. 1.3 Model organisms In the middle of the 19th century, Gregor Mendel conducted foundational research on inheritance using pea plants and ultimately initiated the science of genetics. His experiments also outlined the criteria by which an organism is chosen for research. The selection of the model organism is based on its features and evolutionary relationship to the group of interest (e.g., eukaryotes) and the ability to conduct experiments using them (Müller and Grossniklaus, 2010). Model organisms have become essential tools in gene function research, the effect of different medicines or treatments, and studying physiology and development. Current day model species share several characteristics: the ability to grow in culture, fast proliferation, fully sequenced genome, and amenability to genetic manipulation. There are representative organisms for each broad phylogenetic group (Müller and Grossniklaus, 2010). Prominent examples include Escherichia coli, representing Gram-negative prokaryotes, and brewer’s yeast (Saccharomyces cerevisiae), representing yeasts and other eukaryotes. Invertebrate animals are often represented by the nematode Caenorhabditis elegans and fruit fly (Drosophila melanogaster). Zebrafish (Danio rerio), African clawed frog (Xenopus laevis), and house mouse (Mus musculus) represent vertebrate animals. Multiple model organisms 21 are used for plant research. This section will delineate them, including those selected for experiments in this thesis, P. patens and hybrid poplar P39 (Populus alba x Populus grandidentata). 1.3.1 Plant model systems A representative species from nearly every major plant group enables the study of the plant kingdom and allows for comparison across the plant lineage. Due to recent advances in genome and transcriptome sequencing, numerous plants are now suitable for a variety of research interests such as genetics, biochemistry, and physiology (Koornneef and Meinke, 2010). Thale cress (A. thaliana) is an annual flowering plant of the Brassicaceae family and is the most researched plant model organism (Koornneef and Meinke, 2010). A. thaliana was initially studied in the 1940s, where researchers noted that the plant would be suitable as a model organism following karyotyping experiments that revealed smaller chromosomes than most plants (Koornneef and Meinke, 2010). However, plant scientists of the time focused on plants grown in cell culture (e.g., tobacco species). A resurgence of Arabidopsis research occurred in the 1980s after the discovery of its small genome size, the exchange of research at conferences, and the funding by agencies at the time (Koornneef and Meinke, 2010). A. thaliana exhibits the standard prerequisites of a model organism, such as fast growth and short life cycle, but it also self-pollinates, produces many seeds, and is transformed readily. Research using the model plant organism A. thaliana is valuable. However, it best represents smaller-sized, annual flowering plants. In addition, utility from the model is limited when there is interest in investigating traits of other prominent plant groups, such as woody or non-vascular plants, or when the gene-of-interest is not present in the A. thaliana genome as with 22 EG16. In such cases, other plant model organisms can be utilized. Examples will be presented below. Chlamydomonas reinhardtii is a species of unicellular flagellate chlorophyte green algae (Harris, 2001). The species is often used for studying photosynthesis and motility. C. reinhardtii is readily grown in a lab setting without necessitating vitamins or special additives, and it can grow in light-free environments with the supplementation of acetate. C. reinhardtii cells proliferate with a doubling time of 6-8 hours that enable more convenient experimentation (Harris, 2001). Common liverwort (Marchantia polymorpha) is a bryophyte of the liverwort division that has been a model organism since the 19th century in plant anatomy and physiology research (Shimamura, 2016). As liverworts are distantly related to angiosperm plant species, M. polymorpha is highly suited for studying evolutionary questions. Purple false brume (Brachypodium distachyon) is an angiosperm monocot of the grasses (Poaceae) family. Poaceae form an indispensable plant group to study, as human society heavily relies on nutrition from members of this group. B. distachyon have a recent common ancestor with crops such as wheat (Triticum aestivum) and are therefore representative (Vogel et al., 2010). Nicotiana is a genus of angiosperms of the nightshade (Solanaceae) family. Nicotiana benthamiana is an Australian species and is often used in plant-pathogen research due to the plant’s susceptibility to pathogens (Pombo et al., 2020). Nicotiana tabacum is regularly used in molecular biology research as it can be transformed by a wide variety of methods, including electroporation and direct DNA uptake (Ganapathi et al., 2004). Nicotiana plants are often used to produce pharmaceuticals, vaccines, enzymes and antibodies, such as human interferon beta for hepatitis B and C treatment and influenza vaccine components (Ganapathi et al., 2004). 23 1.3.1.1 Physcomitrella patens The common ancestor of tracheophytes (vascular plants) and bryophytes, which comprise mosses, liverworts, and hornworts, existed approximately 500 million years ago (mya) (Morris et al., 2018). Over this time, many differences accumulated between these groups. Tracheophyte tissues are differentiated into more cell types, particularly the highly specialized liquid-conducting vascular tissues that enable considerable vertical growth. In contrast, as bryophytes do not have these tissues, they are limited in size (Evert and Eichhorn, 2013) (Figure 1.9). Despite these differences, the two groups also share a lot of similarities (Cove, 2005). Both have an alteration of generations, photosynthetic tissues, and sexual and asexual reproduction capability. In addition, similar to tracheophytes, the primary cell wall of bryophytes contains cellulose, hemicellulose, pectin, and many of the CAZymes responsible for their generation (Popper and Fry, 2003) (Figure 1.2). P. patens, a true moss (Bryopsida), serves as a model organism for non-vascular plants. It was initially collected and cultured in the 1960s, and there is now a fully sequenced P. patens genome and established molecular tools for genetic manipulation (Rensing et al., 2020). As P. patens is phylogenetically distant from angiosperms, it serves as an excellent model in evolutionary biology. The life cycle of mosses, including P. patens, initially involves the germination of a haploid spore and the growth of the protonema (primarily chloronema-type), which has a high density of plastids (Rensing et al., 2020) (Figure 1.9). Later, the caulonema develops, a cell type that elongates three times faster than chloronema, and from this tissue, perpendicular cellular divisions occur to form into a leafy bud. This structure eventually matures to generate the adult plant, a leafy gametophore with a stem, phyllids, and rhizoids, that produces the sexual organs (Roberts et al., 24 2012) (Figure 1.9). In older plants, the stem is composed of an epidermal layer of small cells with thick walls, thinner-walled parenchyma cells, and water-conducting cells known as hydroids. The phyllids are one cell layer thick except for the midrib and margins (Roberts et al., 2012). Figure 1.9. The life cycle of the moss Physcomitrella patens (reused with permission from (Rensing et al., 2020)). Haploid protonema grows from spore germination. From this tissue, the leafy gametophore develops, and gametangiogenesis produces female archegonium and male antheridium. Fertilization of the egg cell by the sperm brings about the diploid zygote that develops into the sporophyte that produces haploid spores. 1.3.1.2 Populus spp. Populus spp. are woody, deciduous, perennial flowering plants of the family Salicaceae, distributed in the northern hemisphere. Within angiosperm phylogeny, Populus and Arabidopsis are relatively closely related. Both genera belong to the eurosid clade of dicotyledons, which diverged less than 100 mya (Jansson and Douglas, 2007). Despite their evolutionary relationship, Populus is indispensable as an additional model organism for tree species. The study of the cyclical nature of flowering and autumn senescence is inadequate in Arabidopsis as it is an annual 25 flowering plant. In addition, wood and bark development are not properly studied in other model plants, such as seasonal secondary growth (Jansson and Douglas, 2007). Woody plants are challenging to study due to their slow growth, long life cycle, and larger size. However, Populus spp. are well suited as model trees for many reasons, including their relatively faster growth, small genome size, and the comparative ease of genetic manipulation (Tuskan et al., 2006). The trees have also been used in diverse applications such as carbon sequestration, phytoremediation, and wood and paper industries (Isebrands and Richardson, 2014). Populus trees have a long juvenile phase, with up to 10 years of vegetative growth necessary before flowering (Hsu et al., 2006). Populus can be grown in sterile conditions in small containers in a lab setting and cultured by transferring single internodes into a fresh growth medium. The plants can also be grown in a greenhouse to achieve fast growth with the development of woody tissue. While numerous Populus species are used in research, a notable species is black cottonwood (P. trichocarpa), which is the first tree genome to be fully sequenced (Tuskan et al., 2006). Despite this, other species and hybrids are employed in laboratory research due to their faster growth rate (Cronk, 2005). Noteworthy examples are Populus tremula x Populus tremuloides and P. grandidentata x P. alba (P39) (Figure 1.10), which achieve faster growth rates than other Populus species (Tullus et al., 2011). 26 Figure 1.10. Populus alba x Populus grandidentata (P39) hybrid poplar plant. 4.5 months old greenhouse-grown poplar. Scale bar represents 10 cm. 1.4 Functional characterization of plant genes and gene products Characterization of genes and gene products aid in our understanding of gene function and enhances our knowledge of plant physiology and growth. A standard method for elucidating gene functions is through forward genetics (Ahringer, 2006). This approach typically involves identifying and characterizing a genetic cause of a phenotype, often found as a result of screening of mutants. In contrast, a reverse genetics approach describes the characterization of a specific gene and gene product at the initiation of the study, where the mutant phenotype is used to determine gene function (Ahringer, 2006). As sequence information grows, it is easier to identify genes of interest and characterize them through reverse genetics. The subsections below describe the methodology and techniques involved in gene characterization utilizing the reverse genetics process. 27 1.4.1 Bioinformatics Bioinformatics is a field that assists in the analysis and interpretation of biological data (Luscombe et al., 2001). Bioinformatics tools aid in the exploration of large databases and are important to most life science fields, prominently for sequence assembly, genome annotation, gene identification, and gauging protein structure. With significant advancements in the sequencing of genetic material over the past several years, information is more widely available and accessible, especially data from newly sequenced organism genomes and transcriptomes (e.g., 1000 Plant Genome Project) (Matasci et al., 2014). These data are of significant value for the identification of putative genes across the plant lineages. Determining the presence of a gene in a plant genome is also facilitated by bioinformatics. Basic Local Alignment Search Tool (BLAST) is an essential algorithm that searches through a database using a nucleotide or amino-acid sequence query to find similar sequences (Johnson et al., 2008). The resulting sequences can then be used for further analysis, such as determining the presence of a gene in a genome, probing the sequence variability across different groups of organisms, and quantifying the number of orthologs of a specific gene in a genome. 1.4.1.1 Phylogenetics Phylogenetics is the study of the evolutionary relationships between organisms or taxa (Wiley and Lieberman, 2011). Phylogenetic trees are diagrammatical representations of the end product of phylogenetic analysis. These trees serve to infer evolutionary relationships, and assist in understanding the development and origins of the taxa in question (Yang and Rannala, 2012). There are many approaches to modelling phylogenies, such as neighbor joining (NJ), maximum likelihood (ML), and Bayesian inference (BI) (Yang and Rannala, 2012). NJ trees are calculated by a distance-based approach (Saitou and Nei, 1987). A distance matrix is first 28 calculated based on sequence comparison, and then the taxa are grouped according to these distances to produce a tree with a minimum total branch length. NJ is computationally efficient, so the approach is advantageous for calculating trees for large alignments. However, the method is not suitable for highly divergent sequence alignments due to its representation of sequences as distances and thus produces a less reliable tree (Yang and Rannala, 2012). ML and BI are methods that compare each character (DNA nucleotides or amino acids) in an alignment with each other. ML requires a substitution model, e.g., JTT (Jones et al., 1992), which attempts to infer the substitution rate of nucleotides or amino acids in a sequence. Alternate trees are evaluated to determine whether they can be explained by the given model. The result of ML is the tree with the highest probability of explaining the data (Felsenstein, 1981). However, ML is computationally intensive. BI is similar to ML in that the method is also probabilistic, model-based, and works by evaluating the posterior probability that the tree is accurate, based on the data and the model (Rannala and Yang, 1996). However, compared to the ML approach, a major disadvantage of using BI is its tendency to produce excessively high probabilities. These powerful techniques are all useful depending on the dataset that is being evaluated. 1.4.2 In vitro characterization In vitro characterization involves experimentation and research outside of a living organism. Although these types of experiments do not represent the native environment of activity, they offer a highly controlled setting and allow for the scrutiny of aspects that could be challenging to measure in vivo or in situ (within living systems). The observations from in vitro experiments assist in the prediction of how the components behave in their natural state. 29 1.4.2.1 Heterologous production of proteins In vitro characterizations of enzymes involve the heterologous production and purification of the protein of interest. E. coli or yeasts such as Pichia pastoris are often used for such purposes due to their fast growth, ability to produce various proteins, convenient protein recovery, and established protocols and technical resources. However, the failure rate of these systems is not insignificant, in part, due to folding and stability issues leading to aggregation and missing/unknown co-factors leading to inactivity (Nelson and Cox, 2004). In this case, other expression hosts can be utilized, such as insect, mammalian, or plant cells, which can better accommodate glycosylated proteins (Desai et al., 2010). Additionally, a widely used technique includes transient expression in different organisms. For plants, Nicotiana spp. are commonly used. However, this procedure is more suited for the small-scale production of proteins (Desai et al., 2010). V. vinifera and P. trichocarpa EG16 proteins were produced recombinantly in E. coli cells (Eklöf et al., 2013, McGregor et al., 2017). Those studies provided a foundation on which additional EG16 proteins could be produced similarly and be functional in activity assays. 1.4.2.2 Enzyme activity and substrate specificity of glycoside hydrolases Enzymes are categorized into six functional classes based on the reaction they catalyze. Oxidoreductases operate by transferring electrons between molecules, transferases transfer functional groups from one molecule to another, and hydrolases cleave chemical bonds using water. Lyases modify double bonds, isomerases alter isomeric forms of molecules, and ligases catalyze the formation of chemical bonds with energy, typically from the cleavage of ATP (Nelson and Cox, 2004). As the work in this thesis involves research on a glycoside hydrolase (EC 3.2.1), the following two sections will discuss the characterization process of such enzymes. 30 There are several ways of measuring glycoside hydrolase enzyme activity. The most common ones are based on the quantification of reducing sugars. Reducing sugars are monosaccharides and saccharide units at the ends of di-, oligo- and polysaccharides with a free anomeric carbon/hemiacetal group. The reducing sugar is called so because of its characteristic as a reducing agent toward oxidizing metal salts (e.g., Cu2+). Fehling’s qualitative assay, Tollens’ test and copper-bicinchoninic acid (BCA) assay are colorimetric methods that quantify reducing ends (Arnal et al., 2017). These tests can determine the activity of glycoside hydrolases, which generate reducing ends through hydrolysis of glycosidic bonds. Specifically, the BCA assay, which was effectively utilized in the characterization of VvEG16 (McGregor et al., 2017), works by employing copper ions which are reduced from Cu2+ to Cu1+ ions by the reducing end of sugars. The Cu1+ ion and bicinchoninic acid form a purple complex that can be quantified by spectrophotometry at 562 nm. Using the BCA assay, substrate specificity and enzyme rate constants can be measured. Assays can be carried out using a variety of potential substrates to narrow down a preference. Enzyme rates can be further probed with positive substrates. Michaelis-Menten kinetics is the most widely used model for calculating enzyme rate constants (Michaelis and Menten, 1913, Cornish-Bowden, 2013). The model applies to a system where an enzyme (E) binds the substrate (S) reversibly to form an enzyme-substrate complex (ES) and generates product (P) and the free enzyme (Figure 1.11A). The Michaelis-Menten model relates the maximum reaction velocity (Vmax) at saturating substrate concentration, the rate constant KM, which describes the substrate concentration at 50% of Vmax and the concentration of substrate [S] (Figure 1.11B). The rate constants calculated by the model provide insights into the activity of the enzyme. Specifically, the turnover number, kcat, describes the number of conversions from the substrate to the product 31 per unit of time. Additionally, KM can be defined generally as a measure of substrate affinity under standard conditions. The catalytic efficiency (expressed as kcat/KM) describes the efficiency of the enzyme in converting the substrate into a product. Utilizing this model, previous experiments using VvEG16, showed that the enzyme is more efficient by about an order of magnitude on MLG as a substrate (7.55 mL·mg−1 s−1) than on xyloglucan (0.56 mL·mg−1 s−1) (McGregor et al., 2017). Figure 1.11. Michaelis-Menten model. (A). The model pertains to a system where the enzyme [E] reversibly binds to the substrate [S], producing product [P], and releasing free enzyme. (B). The Michaelis-Menten equation. 1.4.2.3 Enzyme product analysis Product analysis of enzymes is the determination of the specific digest products following enzymatic activity. Two standard methods for analyzing carbohydrate products are high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD) and matrix-assisted laser desorption ionization coupled to time-of-flight mass spectrometry (MALDI-TOF). HPAEC-PAD is a liquid chromatography method that separates and detects molecules in a given sample, especially carbohydrates, such as mono- and oligosaccharides (Rohrer et al., 2013). The process involves the injection of the sample into a column containing a positively charged stationary phase and a strongly alkaline, mobile, aqueous phase. As the carbohydrates in the sample pass through the column, they are separated based on their pKa and three-dimensional structure. The molecules in the sample oxidize when they reach the detector. 32 The amount of sample in the fraction is quantified by charge and displayed as a chromatographic peak. For identification, the retention of a given peak is compared to that of a known standard. MALDI-TOF is a technique frequently used to analyze the mass of biomolecules, such as peptides and carbohydrates (Jurinke et al., 2004). This method relies on the ionization of the analyte through laser irridation of a co-precipitated spot of sample and matrix compound (typically small aromatic acids). The ionized molecules are then accelerated into the mass analyzer, which measures the mass by the time to reach the detector. Using the above techniques, product analysis of VvEG16 and PtEG16 enzymes showed that they cleave MLG at the β(1,4) linkage before the β(1,3), producing glucose (G), cellobiose (GG), and the mixed-linkage tetrasaccharide G3GGG [β-D-Glcp-(1,3)-β-D-Glcp-(1,4)-β-D-Glcp-(1,4)-β-D-Glcp] (Figure 1.12) (Eklöf et al., 2013, McGregor et al., 2017). This site of hydrolysis is unique compared to other MLG-active endo-glucanases (Figure 1.12). In contrast, VvEG16 and PtEG16 cleave xyloglucan after the unbranched glucosyl residue in the backbone (Figure 1.3), as is expected of other xyloglucan endohydrolases (Eklöf et al., 2013, Kaewthai et al., 2013, McGregor et al., 2017). 33 Figure 1.12. MLG hydrolysis site of VvEG16 and other MLG endo-glucanases (reused from (McGregor et al., 2017)). Solid black arrows indicate the primary cleavage site, whereas dashed black arrows indicate the site of secondary activity. 1.4.3 In planta and in situ characterization In situ characterization describe methodologies that are performed “on site” of the object of study. In planta refers to the experimentation carried out in plant tissues specifically. These experiments have many advantages as they give additional insight into the gene or gene product environment. Below are sections that describe techniques that are used for in planta characterization of genes and their products. 1.4.3.1 Transgenic techniques Direct genetic manipulation of plant genomes was first carried out in 1984 in a Nicotiana spp. by utilizing the bacterial pathogen Agrobacterium tumefaciens (Birch, 1997). Since then, many more plants have been added to this list. These techniques form the cornerstone of studying the function of a specific part of the genome. Inhibition, deletion, or addition of genetic material can assist in the determination of their effect on plant physiology and allow for a more robust assessment of their function. 34 There are a variety of different techniques that are commonly used for genetic engineering in plants. A. tumefaciens is a bacterial plant pathogen that aids in plant transformation. As part of the life cycle of this soil bacterium, it integrates genetic material into a plant host genome that induces the production of opines and phytohormones, producing a tumour in the plant (Klee et al., 1987). The bacteria, in turn, use these products as a carbon source of nutrition. Utilizing the bacterial mechanism, virulent genes, specific genetic material, and a resistance gene can be inserted and selected for using reagents such as antibiotics. This technique is widely used in mostly angiosperm plants (Keshavareddy et al., 2018). Another technique for plant transformation is the transfer of genetic material into protoplasts by lipofection (Keshavareddy et al., 2018). This method may be utilized for plants less amenable to Agrobacterium-mediated transformation. Lipofection, specifically polyethylene glycol (PEG)-mediated transformation, is widely used to transform P. patens protoplasts (Liu and Vidali, 2011). This method relies on the DNA repair mechanism of the plant to integrate the recombinant DNA by the direction of flanking regions that are designed to match a specific section of the genome. 1.4.3.2 Phenotype analysis The phenotyping process refers to the method of quantifying physiological traits and the biochemical makeup of organisms (Walter et al., 2015). Phenotyping is often used to determine whether genetic manipulation (e.g., gene deletion) affects a specific trait or biochemical property of the organism. Phenotyping of transgenic plants initially involves assessing the whole organism with subsequent emphasis on the anatomical parts or physiological processes suspected of being affected by the manipulated gene. The expression pattern of the gene would direct phenotyping to 35 that specific tissue. Involvement in a biochemical pathway would lead one to quantify the products of the enzymes in this pathway. Differences to the wild-type in a transgenic plant may indicate that the genetic change led to the observed phenotypes. However, since plant systems are complex, it is not straightforward to conclude that the two systems being compared are precisely the same save for the genetic manipulation. Change in genes may be complicated by indirect effects to the plant biochemistry, such as gene expression differences resulting from physiological changes (Ahanger et al., 2017). Therefore, plant phenotyping is often the first step to identify a more direct mechanism that could explain the traits or processes observed in the transgenic plants. 1.4.3.3 Gene expression analysis Gene expression is a process by which DNA is transcribed to generate non-coding RNA or mRNA which is translated to produce protein. Expression pattern describes the level of expression of a specific gene in certain tissues and cell types of an organism. Determining the expression pattern can help understand when in the plant life cycle and in which tissues the gene product is needed to fulfill its biological function. Determining expression can be done with various methods, including but not limited to Northern blotting, DNA microarray, RNA-sequencing (RNA-seq), and reverse transcriptase quantitative polymerase chain reaction (RT-qPCR). Northern blotting and RT-qPCR aim to quantify the expression of a single gene or a small number of genes at a time. Northern blotting is a technique that involves the immobilization and separation of RNA onto a membrane followed by probing with a labelled complementary DNA or RNA sequence. However, the technique is prone to RNA degradation, and low starting material can result in a false-positive result. RT-qPCR involves making complementary DNA (cDNA) from an RNA sample by reverse transcriptase and 36 then tracking the amplification of specific genetic material using specific primers and fluorescent DNA probes that bind to double-stranded DNA. The technique is widely used as it is sensitive and specific. DNA microarray and RNA-seq are genome-wide approaches to determining gene expression. DNA microarrays is a technique that uses a DNA chip containing DNA sequence probes that hybridize to cDNA of a sample that can be detected and quantified. RNA-seq involves the use of next-generation sequencing of cDNA, which produces a transcriptome. RNA-seq has many advantages over other techniques; it has a wide range of detection, prior knowledge of the genome is unnecessary, and it provides a high sequence resolution (Ozsolak and Milos, 2011). There are also a variety of ways of determining the expression profile in planta, such as in situ PCR, in situ hybridization (ISH), and gene promoter-indicator fusions (such as β-glucuronidase (GUS)) (Athman et al., 2014). In situ PCR and ISH are methods that do not involve plant transformation and can determine expression at a single-cell level. In situ PCR consists of a PCR performed on intact tissues, typically sectioned material, using labelled nucleotides, antibodies directed against those nucleotides and their subsequent detection by microscopy (Athman et al., 2014). In comparison, ISH is carried out similarly but without the amplification of the target gene. Consequently, probing genes with low transcript abundance can result in a false positive (Athman et al., 2014). Promoter-β-glucuronidase (promoter-GUS) analysis involves transforming plants with the promoter of the gene-of-interest followed by the uidA gene from E. coli that encodes for the GUS protein, by itself or fused to the gene-of-interest (Jefferson et al., 1987). When the transgenic plants are treated with a specific substrate (e.g., X-gluc), the GUS protein will convert it into a visible blue product that can be viewed in the plant tissues and acts as a reporter for promoter activity. 37 1.4.3.4 Protein localization Protein localization is the migration of proteins to their functional environment in the cytosol, organelle, or cell wall. Determining the localization of the protein of interest is essential to understanding its function, as it informs the context of the protein activity and its potential interactions (Goodin, 2018). As described above, EG16 localization is unknown. Due to the conflicting nature of the predicted cytosolic localization and location of their in vitro demonstrated cell wall substrates, this analysis is critical in the characterization of EG16. Signal peptides are determinants of subcellular localization (Owji et al., 2018). Proteins targeted to the endomembrane system have N-terminus signal sequences that direct them to the secretory pathway. Subcellular localization prediction algorithms rely on these sequences or lack thereof (e.g., SignalP) (Almagro Armenteros et al., 2019). However, localization by experimental means is still performed to confirm these predictions and gain additional information, such as localization to specific areas of the cytosol. A standard technique for identifying protein subcellular localization involves observing the recombinant protein tagged with a fluorescent protein in the plant tissues using microscopy. This technique is advantageous because the recombinant protein can be viewed in vivo by live imaging, but the process requires cloning and plant transformation. In addition, there are potential downfalls to using this technique. Depending on the promoter, it is possible to obtain low gene expression, or the protein could be mislocalized due to overexpression (Lisenbee et al., 2003). The gene could also be silenced, a high amount of the protein could cause cellular stress, or autofluorescence may obscure the protein localization (Billinton and Knight, 2001). Finally, the addition of the fluorescent protein could cause aggregation or misfolding and, therefore, mislocalization (Wiedenmann et al., 2009). Another way of determining subcellular localization is by 38 immunofluorescence microscopy using antibodies. Antibodies can be made for a wild-type protein or directed against a molecular tag fused to the gene of interest. Advantages of this technique include the highly sensitive nature of antibodies and the ability to stain multiple elements simultaneously. There are many limitations to this method; antibodies are expensive, take a long time to make, and animals are used in their preparation. In addition, the antibody may be nonspecific, and therefore experiments using them require many controls. Moreover, antibodies have to be used on fixed tissue, which could introduce artifacts resulting from the fixation process (Zheng et al., 2017). 1.5 Research aims As the EG16 clade was discovered recently, their sequence diversity across the plant lineages, whether their activity is conserved, and their biological function are undetermined. The current understanding of EG16 function and structure is based on the characterization of orthologs from two angiosperm species and is therefore limited. The biological function of EG16 is unknown and difficult to predict due to the contrast between the expected localization and EG16 substrate availability. Although EG16 gene product function may be vestigial, the aforementioned relatively high expression in P. patens and presence in diverse plant clades suggest otherwise. The primary aim of this thesis is to elucidate the diversity, biochemistry and biological role of EG16 enzymes in the context of plant evolution. This aim will be achieved by identifying EG16 orthologs across eukaryote and plant genomes and transcriptomes via bioinformatic analysis and enzyme activity experiments. In addition, this thesis will present expression tests, localization studies, and phenotyping studies of EG16-downregulated plants of P. patens and hybrid poplar P. alba x P. grandidentata. 39 The work in chapter 2 aims to investigate the evolutionary origin, distribution, and frequency of EG16 genes in the plant kingdom and other eukaryotes using bioinformatic analyses. Prior to the initiation of this project, knowledge of EG16 gene diversity was based on only 33 putative EG16 orthologs due to the limited number of plant genomes and transcriptomes at the time (Eklöf et al., 2013, McGregor et al., 2017). This analysis sought to illustrate the degree of conservation of EG16 orthologs, their frequency in individual genomes, and their distribution among the major plant clades. Most importantly, this analysis aimed to discern whether the hypothesis that EG16-like genes predate XTH genes in the plant lineage is supported. Genomic and transcriptomic databases were utilized to collect EG16 and XTH genes from plant and marine eukaryotes. Maximum-likelihood phylogeny was estimated from the several hundred collected orthologs. Characterization of the enzymatic activity and structure of the bryophyte P. patens EG16 is described in chapter 3. The intent is to discern the substrate specificity, activity, hydrolysis products, and enzyme structure, as well as to compare these aspects to previously characterized EG16 gene products. To this end, PpEG16 was produced in a heterologous manner in E. coli, purified, and biochemically characterized using various techniques. BCA assay was utilized for determining the activity and substrate specificity, while HPAEC-PAD and MALDI-TOF experiments were performed for enzyme product analysis. In addition, ex vivo biomechanical assays using PpEG16 and previously characterized EG16 gene products were performed on etiolated hypocotyls to determine their effect on plant tissues. Lastly, chapters 4 and 5 aim to investigate the biological function of EG16 in P. patens and Populus, respectively. These projects aim to determine the gene expression pattern, protein subcellular localization, and the growth phenotypes of plants downregulated for EG16 expression. 40 To these ends, immunolocalization was performed to determine subcellular localization, and EG16-downregulated growth was compared to wild-type plants. Furthermore, the expression was probed in transgenic Populus plants expressing EG16 promoter::GUS. 1.6 Significance The projects within this thesis uncover important aspects of the EG16 clade. The bioinformatics research showcase the ancient origins of EG16 genes in green algae. As a whole, this thesis highlights the conservation of EG16 homologs in their sequence, enzymatic activity, and biological function across diverse plant lineages. This work is the first to explore the biological role of the EG16s, which identified them as cell wall enzymes associated with young, growing tissue. 41 Chapter 2: Comprehensive cross-genome survey and phylogeny of glycoside hydrolase family 16 members reveals the evolutionary origin of EG16 and XTH proteins in plant lineages 2.1 Introduction Carbohydrate‐active enzymes (CAZymes) are essential and abundant in plant proteomes (Ekstrom et al., 2014). CAZymes perform the synthesis and modification of the diverse polysaccharides that constitute the plant cell wall and are also involved in pathways related to signalling, defence, and energy storage (Henrissat et al., 2001). The actively curated CAZy database (http://www.cazy.org) categorizes CAZymes (namely, glycosyltransferases, glycoside hydrolases, polysaccharide lyases, carbohydrate esterases and auxiliary activities) into families based on protein sequence similarity, which reflects evolutionary history, tertiary structure, and catalytic mechanism (Lombard et al., 2014). The glycoside hydrolases (GHs) form the largest group (presently 171 families) and catalyze the hydrolysis or transglycosylation of glycosidic linkages (Sinnott, 1990, Gilbert et al., 2008). Of the many GH families found in plants, glycoside hydrolase family 16 (GH16) is of particular relevance because it includes xyloglucan endotransglycosylases (XETs, EC 2.4.1.207) and xyloglucan endohydrolases (XEHs, EC 3.2.1.151), encoded by the xyloglucan endotransglycosylase/hydrolase (XTH) gene family. These enzymes have been implicated in the remodelling of the ubiquitous streptophyte, cell wall matrix glycan, and sometimes seed storage polysaccharide, xyloglucan (Rose et al., 2002, Popper, 2008, Eklöf and Brumer, 2010). As members of GH16, XETs, and XEHs share sequence and structural similarities, including common catalytic machinery, with bacterial β(1,3)/β(1,4) mixed‐linkage endo‐glucanases (MLGases, also known as licheninases, EC 3.2.1.73). Early phylogenetic analyses have suggested that bacterial 42 MLGases and plant XETs/XEHs share a common ancestor among the diverse endo‐glucanases and endo‐galactanases of GH16 (Figure 2.1; (Barbeyron et al., 1998, Michel et al., 2001). Figure 2.1. Proposed evolution of glycoside hydrolase family 16 (GH 16) enzymes (Michel et al., 2001; Eklöf et al., 2013) (reused from (Behar et al., 2018)). The precise evolutionary trajectories of the bacterial MLGases and plant XETs/XEHs remain unclear because of significant protein structural differences and a correspondingly vast difference in substrate specificities. However, recent work has uncovered a unique group of GH16 endo‐glucanases (EG16s) among select plant genomes, members of which were proposed to represent extant evolutionary intermediates between the bacterial and plant enzymes (Eklöf et al., 2013, McGregor et al., 2017). Characterization of defining members from Populus trichocarpa (PtEG16) (Eklöf et al., 2013) and Vitis vinifera (VvEG16) (McGregor et al., 2017) in vitro revealed that these enzymes are structurally and functionally intermediate between MLGases and XETs, notably possessing a minimal β‐jellyroll structure and catalytic promiscuity for both MLG and 43 xyloglucan hydrolysis. In particular, EG16 members lack both the C‐terminal extension characteristic of XTH gene products (XET_C: InterPro IPR010713, PFAM PF06955) and a key loop that constricts the active‐site cleft of bacterial MLGases (Figure 2.2). Figure 2.2. Comparison of bacterial mixed‐linkage endo‐glucanases, plant endo‐glucanases (EG16), and plant xyloglucan endotransglycosylase/hydrolases (XTHs) from glycoside hydrolase family 16 (GH16) (reused from (Behar et al., 2018)). Three‐dimensional structure representations were generated using data in the corresponding RCSB Protein Data Bank. Secondary structure topology diagrams were produced manually (β‐sheet lengths are proportionate, loops are not). Tabular data are from Eklöf et al. (2013), McGregor et al. (2017) and the present analysis. 44 Foundational enzymology and structural biology studies of PtEG16 and VvEG16 have enabled higher resolution bioinformatics analysis and more careful delineation of EG16 members, some of which were previously annotated as XTH gene products (e.g., (Yokoyama et al., 2010)). EG16 members had been overlooked as a distinct group within the limited number of plant genomes known at the time, likely for two reasons. First, although they are found in several model plants, EG16s are notably absent from many eudicots, including the model species Arabidopsis thaliana (Eklöf et al., 2013, McGregor et al., 2017). Second, in plants that encode an EG16, only a single ortholog is present in the genome, whereas XTH homologs are usually present in groups of 20–60 genes per species (Figure 2.2; (Eklöf and Brumer, 2010)). An additional curiosity is that EG16 members lack apoplastic secretion signal peptides, ubiquitous in XETs and XEHs, thus precluding the definitive attribution of a cell wall modifying role (Eklöf et al., 2013, McGregor et al., 2017). As such, their biological role remains unclear. In conjunction with biochemical and structural analyses, surveys of the limited number of plant genomes available in 2013 and 2016, predominantly of eudicots and grasses, allowed the identification of a total of ~30 EG16s in ~40 genomes (Eklöf et al., 2013, McGregor et al., 2017). The recent, rapid expansion of plant whole‐genome and large‐scale transcriptome sequencing efforts now provides a much richer data set to explore the distribution of EG16 members across plant lineages. In particular, an abundance of sequence information on the major clades of plants and algae outside the angiosperms, and diverse and previously unexplored lineages of flowering plants, now facilitates the exploration of the emergence of EG16s vis‐à‐vis XTH gene products (Goodstein et al., 2012, Vandepoele et al., 2013, Matasci et al., 2014, Proost et al., 2015). Presented here is a comprehensive bioinformatic survey of GH16 members among Viridiplantae 45 genomes and close relatives, which reveals the emergence of EG16 orthologs and the closely related EG16‐2 subfamily in green plants, concordant with the early divergence of these enzymes. 2.2 Materials and methods 2.2.1 Data sources Gymno PLAZA 1.0, Pico‐PLAZA 2.0 (Vandepoele et al., 2013, Proost et al., 2015), JGI Phytozome v12.1 (Goodstein et al., 2012) and NCBI (NCBI Resource Coordinators, 2017) translated genomic databases (with 119 genomes in total and one transcriptome) were downloaded from the official websites during June–August 2017. Translated transcriptomic data (1459 transcriptomes) were accessed from the 1000 Plants (1KP) database between June 2016 and 2017 (Johnson et al., 2012, Matasci et al., 2014, Wickett et al., 2014). Taxonomic data were obtained from the NCBI Taxonomy Browser (Federhen, 2012), accessed from May to October 2017. Quantification of GH16 members was performed using only well‐curated, assembled genomes from Gymno PLAZA 1.0, Pico‐PLAZA 2.0 and Phytozome v12.1, accessed in July 2017. The Physcomitrella patens EG16 sequence (Pp3c21_9420V3.9) from the JGI Phytozome database was used in the present analysis; sequences XP_001783128.1 from NCBI and PP00353G00180 from PLAZA were omitted because the sequence alignments compared with all other EG16s suggested probable sequencing errors or misassembly. Penium margaritacium genomic and transcriptomic data were sequenced as described (Jiao et al., 2020) (deposited in http://bioinfo.bti.cornell.edu/Penium). Chara braunii and Klebsormidium nitens genomes were available from (Nishiyama et al., 2018) and (Hori et al., 2014), respectively. The programming language Perl was used along with the development environment Eclipse for data curation, organization, and sequence handling. Microsoft Office Excel and 46 Notepad++ were also used for the categorization and organization of data and the preparation of FASTA files. Aliview (Larsson, 2014) was used for the visualization and manual adjustment of sequence alignments made with the embedded multiple sequence comparison by log-expectation (MUSCLE; (Edgar, 2004)) distribution (32‐bit version 3.8.425). Tertiary structures were extracted from the Protein Data Bank (PDB) and visualized with Pymol 2.0. Adobe Illustrator CC 2015 was used to assemble all figures. 2.2.2 Identification of EG16s and XTHs All 33 EG16 sequences included in the bioinformatic analysis of McGregor et al., 2017, and 35 representative XTH sequences were aligned using MUSCLE and separated into two FASTA files (Supplementary Material 2.9 and 2.10) to produce sequence logos via WebLogo (Schneider and Stephens, 1990, Crooks et al., 2004), to aid in the subsequent manual identification and differentiation of EG16 and XTH homologs. Five signature sequence regions were identified as a result (Figure 2.4). Based on preliminary analyses, 15 representative query sequences were selected to probe translated genomic and transcriptomic data sets using protein BLAST (Altschul et al., 1990, Johnson et al., 2008). These 15 queries represented EG16s and XTHs from a diversity of plant lineages, and a bacterial MLGase control: Bacillus subtilis BsLicheninase WP_024571825.1; Brachypodium distachyon BdEG16 KQK23978.1; Amborella trichopoda AtEG16 XP_006857906.1; Gossypium arboretum GaEG16 KHG22293.1; Medicago truncatula MtEG16 XP_013444946.1; P. patens PpEG16 XP_001783128.1; P. patens PpXTH1 CAH58715.1; Picea abies PaEG16 PAB00045440; Populus tremula × Populus tremuloides PttXET16A AAN87142.1; P. trichocarpa PtEG16 XP_002301319.1; S. moellendorffii SmEG16 XP_002960511.1; Tropaeolum majus TmNXG1 CAA48324.1; V. vinifera VvEG16 47 XP_002273975.1; Zea mays ZmEG16 NP_001150346.1; Z. mays ZmXTH1 CAI44139.1 (all are GenBank accessions, except P. abies from PLAZA). All sequences resulting from protein BLAST analysis of individual organisms were aligned with the original queries using MUSCLE within AliView, and were manually examined for the presence of the signature motifs before segregation into ‘EG16’, ‘XTH’, or ‘other GH16’ sequence subfamilies (Supplementary Material 2.1). Sequences were also analyzed with NCBI Batch Web Conserved Domain Search (NCBI Resource Coordinators, 2017) to confirm the presence of GH16 conserved domains. The presence of signal peptides was analyzed using SignalP 4.1 (Petersen et al., 2011). 2.2.3 Phylogenetic analysis EG16, EG16‐2, XTH, and MLGase amino acid sequences were aligned using MUSCLE with default settings within AliView. Following manual curation in AliView to remove apparent fragment sequences, manual realignment was performed using tertiary structural information from the following characterized GH16s as a guide: B. subtilis licheninase (PDB ID 3O5S) (Furtado et al., 2011), Paenibacillus macerans licheninase (PDB ID 1U0A) (Gaiser et al., 2006), T. majus NXG1 (PDB ID 2UWA) (Baumann et al., 2007), and V. vinifera EG16 (PDB ID 5SV8) (McGregor et al., 2017) (see figure 1 in (Eklöf et al., 2013)). The N and C termini were trimmed such that the FASTA alignment file used as input for phylogenetic analysis comprised only the core β‐jellyroll domain (Supplementary Material 2.8). A maximum‐likelihood (ML) phylogeny was estimated using RAxML 8.2.10 on the CIPRES Science Gateway v3.1 (Miller et al., 2010), with the following settings: JTT amino acid substitution matrix, 25 distinct rate categories, and rapid bootstrapping with automatic halting (408 rapid bootstraps). The resulting phylogeny is provided 48 in NEXUS format in Supplementary Material 2.7. FigTree was used for visualization (http://tree.bio.ed.ac.uk/software/figtree). 2.2.4 Expression data Expression data for EG16 orthologs in P. trichocarpa and P. patens were retrieved using the electronic fluorescent pictograph (eFP browser) from The Bio‐Analytic Resource (BAR; (Winter et al., 2007)) using the gene IDs Potri.002G153200.1 and Pp1s353_23V6.1, respectively (Figure A.2 and Figure A.4). Expression data for Pinus abies was acquired from The Plant Genome Integrative Explorer Resource (PlantGenie), specifically, ConGenie, using the ID MA_52109g0010 ((Sundell et al., 2015); Figure A.3). 2.2.5 Accession numbers All accession numbers not otherwise indicated in the text are provided in Supplementary Material 2.1. 2.3 Results and discussion 2.3.1 Cross-genomic census of EG16s and XTHs To provide a comprehensive census of plant GH16 homologs across plant lineages, 126 publicly available genomes and 1460 publicly available transcriptomes were exhaustively mined for the presence of EG16s, XTHs, and other GH16 members. This data set represents a phylogenetically diverse range of species, spanning Viridiplantae (e.g., chlorophyte and charophyte green algae) to land plant clades (e.g., core eudicots), and several groups of marine eukaryotes (e.g., Euglenozoa and red algae) (Figure 2.3). Access to the 1KP transcriptome database, in particular, greatly expanded the phylogenetic coverage; the available whole genomes are dominated by angiosperms (Figure 2.3). 49 Despite the advantages of this expanded data set, the use of translated transcriptomic sequences has particular limitations as a result of incompletely assembled sequences (fragments), with the potential to miss homologs that are present but not expressed, or only weakly expressed, at the time or position of sampling. Nonetheless, the identification of even a single EG16 or XTH member can be used to assign the presence of these subfamilies in a species confidently. As such, the present analysis provides a conservative estimate of the presence of these GH16 members in plants and non‐plant eukaryotes. 50 Figure 2.3. Frequency of EG16 and XTH gene products among plants and marine eukaryotes (adapted from (Behar et al., 2018) and (Jiao et al., 2020)). Tabular data indicate the number of translated genomes and transcriptomes in which at least one EG16, EG16‐2, or XTH member was found, indicated as a ratio of the total number of organisms surveyed in each clade (126 genomes and 1460 transcriptomes in total). All sequences are available in individual spreadsheets in Supplementary Material 2.1. ANA-grade angiosperms are defined as the grade comprising Amborellales, Nymphaeales, and Austrobaileyales. SAR refers to the supergroup containing members of stramenopiles (heterokonts), alveolates, and Rhizaria. Organism phylogeny is based on a reference phylogeny (Wickett et al., 2014) and phylogenies found at JGI Phytozome (Goodstein et al., 2012) and PLAZA (Vandepoele et al., 2013, Proost et al., 2015) (branch lengths are not to scale). N.D. – not determined. Using protein BLAST, automated and manually refined sequence alignments, and the identification of signature sequence motifs (Figure 2.4), the number of identified EG16 orthologs across species was greatly expanded compared with earlier analyses. Specifically, the first study to identify EG16s as a distinct subfamily revealed their presence in 15 of 24 species in 2013 (Eklöf et al., 2013), whereas an updated census in 2016 identified 18 more, for a total of 33 EG16s out of ~40 genomes (McGregor et al., 2017). Now, 587 EG16 orthologs have been identified in 126 genomes and 1460 transcriptomes, representing 1296 organisms. In parallel, over 15,280 XTH sequences, 157 EG16‐2 sequences, 54 MLGase‐like GH16 members, and 304 ambiguous GH16 members have been identified. Supplementary Material 2.1 comprises five individual spreadsheets of all the identified sequences in the groups ‘EG16’, ‘EG16‐2’ (see below for a detailed definition), ‘XTH’, ‘Other GH16’ and ‘Ambiguous’ sequences, respectively, which are fully sortable and selectable on the basis of organism taxonomy (specifically at the levels of genus/species, family, order and major clade), source database, sequence type, and other annotations. The present analysis indicates that canonical EG16s and XTHs are found in all major clades of Streptophyta, but are absent in chlorophyte algae, red algae, and glaucophytes in Archaeplastida 51 (Plantae) (Figure 2.3). Notably, we also identified a distinct group of sequences, here named EG16‐2, which are typified by subtle variations of the EG16 consensus motifs (Figure A.1 cf. Figure 2.4) and an N‐terminal extension of ~150 amino acids that may represent a partial duplication of the GH16 β‐jellyroll domain (Supplementary Material 2.2). EG16‐2 members were observed in two chlorophyte green algae, several charophyte green algae and mosses, individual liverworts and lycophytes, and a few leptosporangiate ferns. The following sections focus on notable observations for specific organism clades, following the order of nesting presented in Figure 2.3. A full molecular phylogeny comprising only high‐quality bacterial MLGase, red algal GH16, EG16, EG16‐2, and XTH sequences is also presented below. Figure 2.4. Aligned sequence logos of canonical EG16 and XTH sequences (reused from (Behar et al., 2018)). Populus trichocarpa PtEG16 (XP _002301319.1) was used as the sequence numbering reference for the EG16 logo plots, and Populus tremula × Populus tremuloides PttXET 16A (ABL 75361.1) was used for the XTH logo plots. 52 2.3.2 EG16 and XTH homologs were not observed in the genomes of non‐plant eukaryotes EG16, EG16‐2, and XTH members were not present in genomes and transcriptomes of Chromista, Dinophyta, and Euglenozoa (Figure 2.3). One dinophyte transcriptome, that of the planktonic organism Prorocentrum micans, was found to have a 107 amino acid fragment similar to other GH16 members (Supplementary Material 2.1, ‘Other GH16’ sheet). Protein BLAST analysis using the NCBI non‐redundant database showed high identity (58%) with a small region identified as a Pfam GH16 domain within a putative sulphate transporter, YbaR, from the dinophyte Symbiodinium microadriaticum (OLP83510.1). Another match was made to a putative GH16 β‐glucanase from a bacterial species, Selenomonas ruminantium (SFT81730.1). Although the fragmentary nature of this sequence currently precludes any definitive assignment within GH16, these results suggest that it is not an EG16 ortholog, but rather belongs to another subfamily (Figure 2.5). Likewise, the Heterokonta, a eukaryotic clade related to Dinophyta that includes kelp and unicellular diatoms (Figure 2.3), contained four species with ‘other GH16’ homologs (Figure 2.5). A total of 10 homologs were identified that share the β‐bulge active‐site motif (EXDXXE) that is characteristic of GH16 laminarinases (Figure 2.1). Indeed, when used as queries against the NCBI database, these putative GH16s align best with GH16 laminarinases (Supplementary Material 2.3). The presence of such gene products in Heterokonta is unsurprising, as the environment that these enzymes may be active in is rich in laminarin, which constitutes ~35% of the cell wall dry weight of Heterokonta and other brown algae (Kadam et al., 2015). Curiously, three sequences from the Chromista alga Synura petersenii (DBYD‐2006863, DBYD‐2050606, and DBYD‐2049612) were discovered in the 1KP database that were highly similar to angiosperm XTHs (Supplementary Material 2.1, ‘Ambiguous’ sheet). This was 53 particularly odd, as these were the only XTH sequences found in a non‐plant eukaryote and searching the available Chromista algae genomes in NCBI did not yield any sequence resembling XTHs. A broader search of the complete NCBI database indicated that DBYD‐2006863 is 85% identical to a Populus tremula × Populus tremuloides XTH (AAN87142.1), DBYD‐2050606 is 85% identical to a Nicotiana tabacum XTH (XP_016507784.1), and DBYD‐2049612 is 93% identical to Medicago truncatula XTH (ACJ85454.1). Thus, it is highly likely that these sequences are contaminants, and they were therefore removed from further analysis. Figure 2.5. Frequency of other GH16 gene products among plants and other eukaryotes (adapted from (Behar et al., 2018) and (Jiao et al., 2020)). Tabular data indicate the number of translated genomes and transcriptomes in which at least one GH16 member, not otherwise identified as an EG16 or XTH member, was identified, shown as a ratio of the total number of organisms surveyed in each clade (126 genomes and 1460 transcriptomes in total). All sequences are available in individual spreadsheets in Supplementary Material 2.1 Organism phylogeny is from JGI Phytozome (Goodstein et al., 2012), and PLAZA (Vandepoele et al., 2013, Proost et al., 2015) (branch lengths are not to scale). N.D. – not determined. 54 2.3.3 Red algal species have MLGase‐like sequences EG16 and XTH sequences were not found in Glauocophyta and red algae; however, protein BLAST analysis using translated red algae transcriptomes revealed 16 fragments with 33–52% identity with a Bacillus subtilis (bacterial) MLGase sequence that was used as a query (WP_024571825.1). The fragments lack the signature EG16 and XTH-defined motifs (Figure 2.4), yet all contain the regular β‐sheet active site motif (EXDXE) that is characteristic of MLGases (Supplementary Material 2.4 cf. Figure 2.1). An NCBI conserved domain search further confirmed that all were GH16 members. Despite the fragmentary nature of these sequences, the available data strongly suggest that they represent MLGases, not EG16 or XTH members. 2.3.4 A limited selection of green algae represents the most phylogenetically distant clades from angiosperms with canonical EG16 and XTH homologs The chlorophyte green algae currently provide the earliest evidence of EG16‐2 and XTH genes in Viridiplantae (Figure 2.3). Chlorophyte cell walls are generally known to contain β(1,4)‐glucan (cellulose), β(1,3)‐glucan and β(1,4)‐mannan, xyloglucan, but not MLG or pectins (Popper et al., 2011, Fangel et al., 2012). Correspondingly, canonical EG16s and XTHs were generally not detected in the available ten chlorophyte genomes and 114 transcriptomes. Three possible exceptions were observed: canonical XTH homologs were found in transcriptomes of the chlorophytes Dunaliella salina, Dunaliella tertiolecta, and Pseudoscourfieldia marina. However, since these sequences are highly similar to angiosperm XTHs, they are likely contaminants and were not included in further analysis (Supplementary Material 2.1, ‘Ambiguous’ sheet). Notably, EG16‐2 sequences were found in the transcriptomes of two chlorophyte species: Botryococcus braunii and Interfilum paradoxum (Supplementary Material 2.1, ‘EG16‐2’ sheet; Figure 2.3). These proteins exhibited distinct, minor variations of the signature motifs of canonical 55 EG16s (Figure 2.4 and Figure A.1), yet these sequences more closely resembled EG16s than other GH16s or XTHs. In particular, these sequences had a regular β‐strand active site motif (EXDXE, Figure 2.1), had high percentage identities with other EG16s, and did not have the C‐terminal extension typical of XTHs. Instead, these EG16‐2 sequences were distinguished by lengthy N‐terminal extensions not otherwise observed in EG16 members (Supplementary Material 2.2). GH16 members distinct from EG16, EG16‐2, and XTH sequences were found in four of 10 chlorophyte genomes (Figure 2.5), namely those of Chlamydomonas reinhardtii (one sequence), Chlorella sp. NC64A (six sequences), Micromonas sp. RCC299 (one sequence), and Ostreococcus lucimarinus (one sequence). All of these sequences contained a β‐bulge, clearly placing them out of the MLGase/EG16/XTH group on the basis of this active site motif (Figure 2.1). Notably, ‘other GH16’ sequences were absent in all Embryophyta (0 of 108 genomes, 0 of 953 transcriptomes; Figure 2.5). In addition, four of the seven charophyte genomes contained GH16 sequences, C. braunii (six sequences), K. nitens (six sequences), Chlorokybus atmophyticus (five sequences), and M. viride (one sequence) (Supplementary Material 2.1, ‘Other GH16s’ sheet; Figure 2.5). EG16s and XTHs also occurred in the charophyte green algae. Of the seven available genomes, EG16-2 and XTHs co-occurred in three genomes, P. margaritaceum (29 XTH and 12 EG16-2 sequences), M. endlicherianum (three XTH and three EG16-2 sequences), and S. muscicola (nine XTH and eight EG16-2 sequences) (Figure 2.6). Notably, one of the charophyte genomes, K. nitens, did not contain XTH homologs but possessed EG16-2 homologs (six sequences), providing evidence of EG16 presence in the most distantly related species to land plants (Figure 2.3 and Figure 2.6). 56 Four species across 46 transcriptomes possessed an EG16 homolog, and nine species possessed at least one XTH homolog. Of these, Desmidium aptogonum and Nucleotaenium eifelense possessed both, a co‐occurrence of EG16s and XTHs in the same species of the most distant charophyte organism. Notably, all 11 of these unique charophyte species, and many other charophytes (33 of 46 total transcriptomes), possessed one or more EG16‐2 sequences (Supplementary Material 2.1; Figure 2.3). As observed for the chlorophytes, EG16‐2 sequences are sufficiently distinctive to warrant separation into an independent group, including a lack of signature EG16 motifs (Figure 2.4 and Figure A.1) and the presence of extended N-termini. Closer inspection of the ‘Ambiguous’ plant GH16 sequences, the sequences of which precluded definitive assignment, indicates that an additional 33 charophyte species may possess XTH‐like sequences (Supplementary Material 2.1). Although there is currently no biochemical characterization of any EG16, EG16‐2, XTH, or XTH‐like protein outside angiosperms, the presence of these sequences coincides with the general observation that species possessing both MLG and xyloglucan in their cell walls were present early in the evolution of Streptophyta (e.g.., charophyte algae) (Popper et al., 2011, Fangel et al., 2012). Notably, recent screening studies have revealed XET activity in Chara, Klebsormidium, and Zygnema tissues (Herburger et al., 2018), possibly arising from one of these GH16 members. 2.3.5 EG16 homologs were observed in many hornwort and liverwort transcriptomes Despite an absence of genomic data, transcriptome analysis revealed at least one EG16 and one XTH simultaneously in six of nine hornworts; Phaeoceros carolinianus additionally has an EG16‐2 ortholog (Supplementary Material 2.1; Figure 2.3). Notably, five hornwort species had EG16 orthologs that were likely to be catalytically crippled, because they lacked one or more key catalytic residues (Supplementary Material 2.5; Figure 2.3). It is presently unclear whether this 57 is simply a result of transcriptome sequencing errors; however, such variants are reminiscent of catalytically inactive GH16 members from invertebrates, which instead function in glycan recognition as part of the innate immune responses in the Toll Pathway (Lee et al., 2009). Analysis of the single available liverwort genome of Marchantia polymorpha revealed the presence of one EG16 and 44 XTH homologs. Analysis of 28 liverwort transcriptomes extended the number of species containing at least one EG16 or XTH to 15 and 24, respectively (Figure 2.3 and Figure 2.6). EG16‐2 sequences were only observed in the transcriptomes of the liverwort Noteroclada confluens (Supplementary Material 2.1). 2.3.6 EG16 homologs are widespread in mosses, lycophytes, and ferns The genomes of P. patens and Sphagnum fallax both encoded EG16s and XTHs (Figure 2.3). Enumeration of these proteins indicated that XTHs were present in large families of tens of homologs, consistent with previous studies (Yokoyama et al., 2010), whereas the number of EG16 homologs was restricted to one each in P. patens and S. fallax (Figure 2.6). Indeed, cross‐genome analysis indicates that EG16s usually occur as single orthologs or are entirely absent, which strongly contrasts with the manifold XTH homologs (Figure 2.6). The genomes of both S. fallax and P. patens encoded a single EG16‐2 homolog each (Figure 2.6). In transcriptomes of 41 moss species, 35 (85%) had an EG16 homolog, 21 had an EG16‐2 homolog (51%), and 38 (92%) had at least one XTH homolog. Although these data are limited as a result of sampling considerations (see above), these results provide evidence of the very high penetrance of the EG16 and EG16‐2 members among the mosses. The genomic databases were limited with regard to lycophyte genomes, with only two available, Selaginella kraussiana and S. moellendorffii, of which S. moellendorffi had a known EG16 homolog (McGregor et al., 2017). Of 21 lycophyte transcriptomes, 12 had an EG16 58 homolog (57%), whereas all had XTH homologs (Figure 2.3). Only the transcriptome of Phylloglossum drummondii contained an EG16‐2 sequence. At the time of this analysis, there were no publicly available genomes for fern species (Sessa et al., 2014); however, EG16s, as well as XTHs, are highly prevalent in fern (monilophyte) transcriptomes. All 12 eusporangiate fern transcriptomes had EG16 and XTH homologs, whereas 58 of 66 (87%) leptosporangiate fern transcriptomes had an EG16 homolog versus 60 (90%) with at least one XTH homolog (Figure 2.3). Five leptosporangiate transcriptomes contained EG16‐2 members. EG16‐2 sequences were not observed in the genomes and transcriptomes of any other vascular plants (i.e., none were found in seed plants; Figure 2.3). 2.3.7 EG16 homologs have been widely maintained among gymnosperms Ginkgoales, which contain one extant species, Ginkgo biloba, had a single EG16 and 13 full‐length XTH homologs present in its genome (Figure 2.3 and Figure 2.6); five XTH fragments were also found (Supplementary Material 2.1, ‘XTH’ sheet). Notably, a sequence retrieved from the PLAZA database (GBI00016679) appeared to comprise an N‐terminal fragment of an EG16, which could be completed to derive the full‐length protein by the overlapping of 119 amino acids with 100% identity to a C‐terminal fragment from the 1KP transcriptome database (SGTW‐2037882; Supplementary Material 2.6). Unusually, the complete G. biloba EG16 sequence was found to contain a secretion signal peptide, which is otherwise typically absent among orthologs and the absence of which is a defining feature of EG16s (Figure 2.2); the presence of a signal peptide in this species thus represents an evolutionary novelty. 59 60 Figure 2.6. Quantitative analysis of EG16, XTH, and other GH16 homologs in translated genomes of plants and marine eukaryotes (adapted from (Behar et al., 2018) and (Jiao et al., 2020)). The light‐grey vertical bar serves as a visual guide for one homolog on the x‐axis. Organism phylogeny is based on a study from a recent publication (Wickett et al., 2014), as well as the phylogeny provided by JGI Phytozome (Goodstein et al., 2012) and PLAZA (Vandepoele et al., 2013, Proost et al., 2015) (branch lengths are not to scale). The single available cycad genome that of Cycas micholitzii is strikingly bereft of XTH members (with two full‐length homologs and three fragments) and completely lacks an EG16 ortholog (Supplementary Material 2.1; Figure 2.6). Analysis of an additional three cycad transcriptomes indicated that all had XTHs, whereas an EG16 was found only in Encephalartos barteri (Figure 2.3). Likewise, the single available Gnetales genome, that of Gnetum montanum, encoded only two canonical XTH homologs and lacked an EG16 ortholog altogether (Figure 2.3 and Figure 2.6); however, analysis of three transcriptomes revealed the presence of an EG16 ortholog in Welwitschia mirabilis, whereas XTH homologs were found in all Gnetales data sets (Supplementary Material 2.1; Figure 2.3). All eight available conifer genomes encoded single EG16 orthologs and tens of XTH homologs each (Figure 2.3 and Figure 2.6). Transcriptome analysis of 72 conifers identified EG16 orthologs among an additional 49 species (68%) versus the ubiquitous identification of at least one XTH per species (Figure 2.3). The disparity between the genomic and transcriptomic databases may reflect the limited, specific expression of EG16 genes and/or a predominance of XTH genes (Figure 2.6). Notably, four Pinales EG16 homologs (Picea glauca PGL00023948, Pinus sylvestris PSY00021134, Picea abies PAB00045440, and Acmopyle pancheri HILW‐2113134) were found to have signal peptides (Supplementary Material 2.1, ‘EG16’ sheet). In 61 fact, these proteins, along with the G. biloba ortholog discussed above (GBI00016679), were the only EG16s predicted to contain a signal peptide among the 587 canonical EG16s identified. 2.3.8 EG16 homologs were variably maintained among angiosperms Apart from eudicots and monocots, the angiosperms include the ANA‐grade lineages (Amborellales, Nymphaeales, and Austrobaileyales, comprising Amborella, water lilies and others), magnoliids, and Chloranthales. Genomic resources for these plants are limited: only a single genome was available, that of the lineage comprising the sister group of all other angiosperms, Amborella trichopoda, the single EG16 ortholog of which was previously identified (McGregor et al., 2017). This organism also contains ~20 XTH homologs (Figure 2.3 and Figure 2.6). Analysis of the available transcriptomes revealed that all three of the aforementioned groups had at least one organism with an EG16 homolog, whereas at least one XTH homolog was found in all of the transcriptomes (Supplementary Material 2.1; Figure 2.3). In sharp contrast to other angiosperms, nearly all (21 of 22) commelinid monocot genomes encoded an EG16 ortholog. The single exception was Musa acuminata (Figure 2.3), a banana species. Transcriptomic data allowed the identification of EG16 orthologs in an additional 15 species (Supplementary Material 2.1, ‘EG16’ sheet; Figure 2.3). Among commelinid monocots, XTH were present as approximately 30 homologs per genome and were identified in all but one transcriptome (Figure 2.3 and Figure 2.6), that of Zea nicaraguensis. Genomes of non‐commelinid monocots, represented by Spirodela polyrhiza and Zostera marina, did not encode EG16 orthologs, despite possessing 12 and 18 XTHs, respectively (Figure 2.3 and Figure 2.6). In contrast, EG16 orthologs were found in 42 of 63 transcriptomes; XTH homologs were ubiquitous in these data (Figure 2.3). 62 The non‐core eudicots are represented by a single genome, that of Aquilegia coerulea, which does not encode an EG16 ortholog but encodes 26 XTH homologs (Figure 2.3 and Figure 2.6). In contrast, analysis of 35 non‐core eudicot transcriptomes identified 20 species encoding an EG16 ortholog; at least one XTH was found in all 35 species (Figure 2.3). Four species of the genus Papaver, which contain numerous poppy species and other frost‐tolerant plants, had EG16 orthologs that have an altered active site motif, suggesting that they are catalytically crippled, as was discussed above for some hornwort orthologs (Supplementary Material 2.5). Notably, a number of orthologs were missing both catalytic residues and the ‘helper’ aspartic acid residue. The distribution of EG16 orthologs among the core eudicots was variable. Our original, more limited analyses indicated that EG16 orthologs are not present in several model eudicots, including members of Brassicaceae such as Arabidopsis thaliana (in rosids) and Solanaceae (nightshade) species (in asterids) (Eklöf et al., 2013, McGregor et al., 2017). Of five core eudicot genomes outside the rosids and asterids, four encode an EG16 ortholog, and all had ~30–50 XTH homologs (Figure 2.3 and Figure 2.6). Analysis of 96 transcriptomes enabled the identification of EG16s in an additional 21 species, in which the observation of at least one XTH homolog was essentially ubiquitous (93 of 96 species). In contrast, EG16 orthologs were strikingly rare in asterids. Among the six genomes surveyed (Supplementary Material 2.1), none had EG16s but, in contrast, had tens of XTH homologs (Figure 2.3 and Figure 2.6). Analysis of 222 asterid transcriptomes revealed only 18 species expressing an EG16 ortholog, versus the essentially ubiquitous identification of one or more XTH homologs (220 of 222 species; Supplementary Material 2.1; Figure 2.3). EG16 orthologs were more common in rosids, but yet were present in only half of the available genomes, in which XTH homologs were ubiquitous and abundant (Figure 2.3 and Figure 2.6). Notably, EG16s were not found in many members of the Brassicaceae 63 family, e.g., Brassica nigra (black mustard), Brassica oleracea capitata (cabbage), and the model plant Arabidopsis thaliana (Supplementary Material 2.1). However, they were common in Fabales, such as the legume crops Glycine max (soy) and Cicer arietinum (chickpea). They were also common in Rosales such as Cannabis sativa (hemp), Fragaria vesca (strawberry), and Prunus persica (peach), and in the woody plants of the family Salicaceae, which includes numerous poplar (Populus) and willow (Salix) species. 2.3.9 The molecular phylogeny of EG16 orthologs is generally concordant with contemporary estimates of Viridiplantae evolution The collection of a combined large number of EG16 and EG16‐2 orthologs across a wide diversity of plant lineages provided a unique opportunity to examine the molecular evolution of these proteins within GH16. The calculation of a maximum‐likelihood phylogeny following iterative MUSCLE and structure‐guided manual alignment of the core β‐jellyroll domain of high‐quality, curated sequences revealed that EG16 (n = 363) and EG16‐2 (n = 38) sequences clearly segregated from each other, and from exemplar bacterial MLGases (n = 3), rhodophyte GH16 members (predicted MLGases, n = 3) and XTHs (n = 21), with high bootstrap support (Supplementary Material 2.7; Figure 2.7). The global topology of this phylogeny essentially recapitulated that seen in previous analyses of GH16 ((Barbeyron et al., 1998, Michel et al., 2001, Eklöf et al., 2013); Figure 2.1), with the exception of the emergence of EG16‐2 sequences as a monophyletic group independent of canonical EG16s. As discussed above, the EG16‐2 group was originally identified on the basis of variations on the canonical EG16 signature motifs (Figure 2.4 and Figure A.1), and is composed of sequences from most green‐plant lineages except seed plants (Figure 2.3). Closer inspection of these sequences, including focused alignment with selected canonical EG16s, XTHs and MLGases 64 (Supplementary Material 2.8), revealed clear similarity with the EG16s in particular. Strong bootstrap support for the segregation of EG16 and EG16‐2 sequences highlights their divergence. The molecular phylogenetic estimate within EG16 and EG16‐2 directly parallels our current understanding of organismal phylogeny (Supplementary Material 2.7 and Figure 2.7, cf. Figure 2.3), pointing to its potential utility in estimating plant phylogeny. The restriction of EG16‐2 members to major Streptophyta lineages, including non-vascular land plants, implies a long parallel evolution and maintenance of these orthologs within genomes (with a loss in seed plants). Within the EG16 clade, orthologs are also segregated into clades that generally recapitulate a contemporary model of Viridiplantae evolution ((Wickett et al., 2014); Supplementary Material 2.7 and Figure 2.7, cf. Figure 2.3). For example, EG16 orthologs of land plants are nested among charophyte algae, in accordance with our understanding of land‐plant origins. Likewise, hornwort EG16s are sister to other land plants, whereas liverwort EG16s form a monophyletic group neighbouring those of lycophytes. Fern EG16s appear in several weakly supported positions across the land‐plant backbone, before the divergence of seed plant orthologs, and with a clade of gymnosperms and another of angiosperms, within which major flowering‐plant clades can be identified (Supplementary Material 2.7; Figure 2.7). 65 Figure 2.7. Maximum‐likelihood phylogeny of EG16 members (reused from (Behar et al., 2018)). 337 EG16s, 38 EG16‐2, 28 XTHs (a minimal random selection representing the diversity of plant clades), three red algae putative MLGases, and three bacterial MLGase high‐quality sequences were aligned with MUSCLE and manual structure‐guided refinement (see Supplementary Material 2.4 for all input sequences and accessions) prior to RAxML analysis with 406 rapid bootstraps. The bacterial MLGases, the ancestors of which are proposed to have given rise to XTHs and EG16s (Planas, 2000, Eklöf et al., 2013, McGregor et al., 2017), were used as an outgroup to root the phylogeny. The output file in NEXUS format is provided as Supplementary Material 2.7. 66 Despite the overall concordance of EG16 molecular phylogeny with plant evolutionary models, some notable exceptions are nonetheless observed. Lycophyte EG16s are arranged into two groups, one between hornworts and mosses, and another between mosses and liverworts. As lycophytes are vascular plants, they are generally considered to be sister to a clade comprising ferns and seed plants (the euphyllophytes). Likewise, EG16 orthologs from some eudicots and ‘early’ angiosperms are arranged together with core eudicots (Supplementary Material 2.7; Figure 2.7). The bootstrap support for these unusual arrangements is poor, reflecting the lack of support for some deep relationships in plant phylogeny, which is not unexpected in any broad phylogeny estimated from a single gene (e.g., (Wickett et al., 2014)). Although definitive phylogenetic placements of some taxa are not possible from this gene alone, the retention of a single ortholog across most major lineages of plants nevertheless suggests strong conservation of EG16 structure across the organismal phylogeny, and perhaps the strong conservation of biological function across a very broad diversity of plant lineages. The inclusion of bacterial MLGases (root) and rhodophyte MLGase‐like sequences in the phylogeny highlights the very ancient divergence of EG16/EG16‐2 members and XTHs. Indeed, the comprehensive census of GH16 members among Viridiplantae and related marine eukaryotes failed to reveal any clade that exclusively possessed EG16s, EG16‐2, or XTHs (Figure 2.3). The original hypothesis was that a possible evolutionary pathway to extant plant GH16 members could be: bacterial MLGase → ancestral plant EG16 (via loop deletion) → plant XTHs (via C‐terminal extension and gene duplication) (Eklöf et al., 2013). Such a pathway might reasonably be anticipated from three‐dimensional structural analysis of these proteins ((Eklöf et al., 2013, McGregor et al., 2017) Figure 2.2) and consideration of gene copy numbers in species. The evolution of a comparatively restricted number of XEHs as a result of key active‐site loop 67 extension within the larger group of XETs was previously revealed by combined biochemical, structural, and phylogenetic analyses (Baumann et al., 2007). The observation of EG16-2 homologs in charophyte species, K. nitens in particular, suggests that EG16 genes are ancestral to extant plant GH16 members (Figure 2.6), supporting the proposed evolutionary pathway. The census data additionally indicate that XTHs arose later, likely within the last common ancestor of the Zygnematophyceae and land plants (Figure 2.6). 2.3.10 Observations on the putative biological roles of EG16 and EG16‐2 proteins The physiological role of EG16 and EG16‐2 proteins is presently unknown and mysterious. In contrast to XTH genes, which have undergone the extensive multiplication typical of other cell‐wall CAZymes (Henrissat et al., 2001, Yokoyama et al., 2004, Geisler-Lee et al., 2006, Del Bem and Vincentz, 2010, Eklöf and Brumer, 2010, Yokoyama et al., 2010), EG16 genes are invariably present as single, highly conserved orthologs in individual species, which is more reminiscent of core metabolic enzymes. On the other hand, EG16 orthologs have been maintained in some plant genomes and lost in others, which suggests a degree of dispensability. It is not clear how to reconcile these contrasting observations. The tissue‐specific microarray data available indicate that expression patterns are comparable among distant plant clades. For example, the expression pattern of EG16 in P. trichocarpa, a woody eudicot, mirrors that observed for the ortholog in Pinus abies, a long‐living conifer species (Figure A.2 and Figure A.3): expression is strongest in young tissues and in reproductive tissues (Wilkins et al., 2009). Likewise, in the moss P. patens, there is a strong expression of the EG16 ortholog in protonemata (cells at the earliest stage of the gametophyte life phase) and in the base of young gametophyte leaves (Figure A.4; (Winter et al., 2007, Yokoyama et al., 2010, Ortiz-Ramirez et al., 2016). 68 A lack of signal peptides in all EG16 members, except those from four Pinales and G. biloba, is particularly puzzling in light of their demonstrated activity on substrates found in the primary cell wall, viz. xyloglucan and MLG (Eklöf et al., 2013, McGregor et al., 2017). This may indicate leaderless secretion, which can involve over 50% of the plant secretome in certain conditions (Agrawal et al., 2010, Ruiz-May and Rose, 2013). It is also possible that EG16 acts intracellularly, perhaps in metabolic roles. 2.4 Conclusions In this study, access to expanded plant genomic and transcriptomic resources has enabled unprecedented insight into the early origins of GH16 xyloglucan‐ and MLG‐active enzymes, which are central to the modification of these key cell wall matrix polysaccharides. Extant EG16 members represent the early adoption of the β‐jellyroll protein scaffold from an early, possibly bacterial or ancient eukaryotic, GH16 member, which diversified by loop deletion and extension of the N-terminus (in EG16‐2 members) or C-terminus (in XTH members). In particular, the unambiguous identification here of the EG16‐2 members as a new subfamily will enable refined studies of these distinct plant proteins. The present comprehensive census will underpin future studies to elucidate the physiological role of all plant GH16 members and serve as a roadmap to delineate the closely related EG16, EG16‐2, and XTH gene products in forthcoming bioinformatics analyses of emerging genomes and transcriptomes (e.g., (Cheng et al., 2018)). Specifically, the large and carefully curated compilation of sequences provided here (Supplementary Material 2.1) may be used to generate hidden Markov models for rapid CAZyme analysis (Lombard et al., 2014). 69 Chapter 3: Conservation of endo-glucanase 16 (EG16) activity across highly divergent plant lineages 3.1 Introduction Glycoside hydrolase family 16 (GH16) forms an important group of enzymes that hydrolyze either β(1,4) or β(1,3) linkages of glucans or galactans. The substrates of bacterial GH16 members are diverse and include agar, porphyran, carrageenan, laminarin, mixed-linkage β-glucan, and xyloglucan, among others (Viborg et al., 2019). In plants, GH16 xyloglucan endo-transglycosylase/hydrolase (XTH) gene products, comprising xyloglucan endotransglycosylases (XETs, EC 2.4.1.207), and xyloglucan endohydrolases (XEHs, EC 3.2.1.151), modify the most abundant matrix glycan in the primary cell wall of most streptophytes (Eklöf and Brumer, 2010). XETs cleave the xyloglucan backbone and subsequently form a new glycosidic bond with another xyloglucan chain, whereas the XEHs irreversibly cleave the glycosidic backbone through hydrolysis (Baumann et al., 2007). Individual XETs and XEHs have been implicated in diverse morphological processes in plants, such as cell growth and expansion, fruit ripening, flower development, and seed germination (Edwards et al., 1986, Rose and Bennett, 1999, Rose et al., 2002, Becnel et al., 2006, Miedes and Lorences, 2009, Eklöf and Brumer, 2010, Kaewthai et al., 2013, Cosgrove, 2016, Stratilová et al., 2020a). Interestingly, some homologs have been shown to catalyze hetero-transglycosylation between xyloglucan and other saccharides (Hrmova et al., 2007, Simmons et al., 2015, Stratilová et al., 2020a, Stratilová et al., 2020b). Recently, refined phylogenetic and biochemical analysis of plant GH16 members uncovered a unique, previously uncharacterized clade comprising endo-glucanase 16 (EG16) orthologs (Eklöf et al., 2013). Two members of this group, Populus trichocarpa EG16 (PtEG16) (Eklöf et al., 2013) and Vitis vinifera EG16 (VvEG16) (McGregor et al., 2017), were characterized 70 as glycoside hydrolases (GH), which cleave both mixed-linkage β(1,3)/β(1,4)-glucan (MLG) and xyloglucan (XyG). The tertiary structure of VvEG16 revealed a broad active site cleft, thus rationalizing the broad substrate specificity of EG16 members. These enzymes also lack the defining C-terminal extension of XETs and XEHs (“XET_C” PFAM domain PF06955 (Johansson et al., 2004, Baumann et al., 2007, Eklöf et al., 2013, McGregor et al. 2017)). Thus, primary and tertiary structural analysis together support that EG16 members constitute a discrete group that appears to be intermediate between extant bacterial MLGases and plant XTH gene products (Eklöf et al., 2013, Behar et al., 2018). Strikingly, whereas plant genomes typically contain 20-60 XTH homologs (Eklöf and Brumer, 2010, Behar et al., 2018, Jiao et al., 2020), it was initially observed that, when present, only a single EG16 ortholog is found (Eklöf et al., 2013, McGregor et al., 2017). In particular, EG16 members are noticeably missing from Arabidopsis thaliana, Nicotiana tabacum, and other eudicots. These observations prompted us to undertake an extensive cross-genome/transcriptome survey of nearly 1300 plant species. It was shown that EG16 protein sequences are both highly conserved and widely distributed across most plant lineages, from non-vascular land plants such as mosses and liverworts to seed plants ferns, gymnosperms, as well as graminaceous monocots and certain eudicots (Behar et al., 2018). EG16 homologs are also found in some green algae (Jiao et al., 2020). The phylogeny of EG16 protein sequences recapitulates contemporary models of plant species evolution, which further reflects a high degree of sequence conservation. The biological function(s) of EG16 members is currently unknown due to a lack of genetic studies. In the present study, we sought to biochemically characterize a representative from a bryophyte plant lineage (Figure 3.1) because only angiosperm EG16s had been characterized previously (Eklöf et al., 2013, McGregor et al., 2017). Thus, we cloned and recombinantly 71 expressed three EG16 variants identified in the transcriptome of the model moss, Physcomitrella patens. Subsequent enzymology defined PpEG16 as an endo-β(1,4)-glucanase/xyloglucanase, with the ability to weaken plant tissues ex vivo. Figure 3.1. EG16 orthologs among land plant clades. (A). Percent similarity (shades of teal) and percent identity (shades of grey) of EG16 protein sequences representing a variety of plant clades. (B). Evolutionary relationships of land plants, based on analysis by Morris et al. 2018. 72 3.2 Materials and methods 3.2.1 Materials All materials and chemicals were purchased from Sigma-Aldrich unless otherwise stated. Phusion High-Fidelity DNA Polymerase, dNTPs, restriction enzymes, and T4 DNA Ligase were purchased from New England Biolabs. Barley β-glucan (MLG; P-BGBH), tamarind xyloglucan (XyG; P-XYGLN), wheat arabinoxylan (wAX; P-WAXYM), curdlan (P-CURDL), konjac glucomannan (kGM; P-GLCML), carob galactomannan (cGM; P-GALMH), and yeast β-glucan (yBG; P-BGYST) were purchased from Megazyme. Carboxymethyl cellulose (CMC; 332641000), hydroxyethyl cellulose (HEC; K391), laminarin (G9752), and starch (S516) were purchased from Acros Organics, Amresco, Millipore-Sigma and Fisher Chemical, respectively. Stock solutions of these polysaccharides were prepared according to manufacturer instructions at 10 mg·mL-1. Azo-dyed and azurine cross-linked (AZCL) xyloglucan I-AZXYG); AZCL-hydroxyethylcellulose (I-AZCEL), and endo-glucanase E-XEGP (Gloster et al., 2007, Park and Cosgrove, 2012) were purchased from Megazyme. P. trichocarpa EG16 (Eklöf et al., 2013) and V. vinifera EG16 (McGregor et al., 2017) were produced essentially as previously described. P. patens EG16 variants were produced as described below. Etiolated hypocotyl walls from chia (Salvia hispanica) and cucumber (Cucumis sativus, cv. Straight Eight, Everwild Farms Inc.) were used for biomechanical assays. 3.2.2 Bioinformatics PpEG16 sequences were extracted from Phytozome: Pp3c21_9420V3.1, Pp3c21_9420V3.9, Pp3c21_9420V3.10 (GenBank accession: XP_024359727.1) (Supplemental 73 Material 3.1). NCBI BLAST was used to analyze the protein sequence identity, and similarity between EG16s from different plant groups (accession numbers are from GenBank unless otherwise stated): P. patens XP_024359727.1; Sphagnum fallax Sphfalx0006s0041.1 (JGI Phytozome); Marchantia polymorpha PTQ31451.1; Selaginella moellendorffii XP_EFJ38050.1; Picea abies PAB00045440 (Plaza); Amborella trichopoda XP_006857906.1; Brachypodium distachyon PNT78322.1; V. vinifera XP_002273975.1; P. trichocarpa XP_002301319.2. Protein sequence alignments were manually refined versus the tertiary structure of VvEG16 (McGregor et al., 2017) and annotated with ESPript (Robert and Gouet, 2014). A structural homology model of PpEG16.9 was generated using the SWISS-MODEL server (Waterhouse et al., 2018). Several models were generated using the following PDB IDs as templates: 5DZE, 5DZF, 5DZG, and 5SV8. The model with the highest Global Model Quality Estimate (GMQE) score, based on PDB ID 5DZG, was selected for analysis (available as Supplemental Material 3.2 and at the Protein Model Database, PMDB ID PM0084148). The overall QMEAN Z-score was 0.75 and QMEANDisCo Global score was 0.83. Model validation by MolProbity revealed statistics comparable to the 5DZG template: MolProbity Score = 1.27, Clash Score = 1.63, Ramachandran Favoured = 94.82 %, Ramachandran Outliers = 1.04%, Rotamer Outliers = 0.59%. Structural alignment of the model with the template was performed in Coot using the SSM (Secondary Structure Matching) method (RMSD 0.438 Å across 160 Cα pairs). Protein representations were produced using PyMOL (Schrödinger, Inc.) 3.2.3 Cloning of PpEG16 targets, transformation, protein production and purification Coding sequences of PpEG16.1, PpEG16.9, and PpEG16.10 were codon-optimized for E. coli expression using the IDT Codon Optimization Tool (Supplemental Material 3.3). cDNA sequences were obtained from Bio Basic (Markham, ON, Canada), each was inserted into pET28b 74 with the restriction sites NdeI and NheI. For crystallography experiments, LIC cloning was utilized for subcloning all targets into pMCSG53 (Table B.1) (Eschenfeldt et al., 2009). PCR reactions were run in Bio-Rad C1000 Thermal Cycler. Constructs were transformed into BL21 (DE3) competent E. coli cells via heat shock. Transformants were used to inoculate 5 mL Studier Zym-505 media (5 g·L-1 yeast extract, 10 g·L-1 tryptone, 25 mM Na2HPO4, 25 mm KH2PO4, 50 mM NH4Cl, 5 mM Na2SO4, 0.5 g·L-1 glucose, 5 g·L-1 glycerol) supplemented with antibiotic (50 µg·mL-1 kanamycin for pET28b constructs or 100 µg·mL-1 ampicillin for pMCSG53 constructs) and grown at 37°C overnight with shaking at 200 RPM (Studier, 2005). Starter culture was added to a 4 L flask with 1 L of media of fresh media and grown until an OD600 value of 1.0 was achieved (measured with an Agilent Cary 60 spectrophotometer). To initiate protein production, 0.2 mM IPTG was added, and the culture was incubated at 16°C overnight shaking at 200 RPM. The culture was centrifuged at 4000 g for 25 min at 4°C, and the resulting bacterial pellet was resuspended (20 mM sodium phosphate buffer, pH 7.5, 300 mM NaCl, 20 mM imidazole, 5 mM DTT, 1 mM EDTA, 2500 units Benzonase Nuclease, 0.06 mg·mL-1 lysozyme, 1x cOmplete™ Protease Inhibitor Cocktail), sonicated (Misonix XL2020 Ultrasonic Liquid Processor) for 3 min at intervals of 20 s on and 20 s off. The sample was then centrifuged at 24,000 g for 40 min at 4°C, and the supernatant was separated from the cell debris. The presence and mass of protein was confirmed with SDS-PAGE (Bio-Rad mini Protean TGX 4–20%) stained with Coomasie blue dye (50% methanol, 40% H2O, 10% acetic acid, 0.25% Coomassie blue R-250). Using a Bio-Rad NGC Chromatography System, the supernatant was loaded on a 1 mL HisTrap affinity column containing nickel-loaded beads (GE Healthcare), which was subsequently washed with 20 column volumes of wash buffer (20 mM sodium phosphate buffer, pH 7.5, 300 75 mM NaCl, 20 mM imidazole, 5 mM DTT, 1 mM EDTA). To elute the protein, the column was washed with a linear gradient of wash buffer to elution buffer (20 mM sodium phosphate buffer pH 7.5, 300 mM NaCl, 400 mM imidazole, 5 mM DTT, 1 mM EDTA) over 20 column volumes. For subsequent size exclusion chromatography (SEC), pooled fractions were concentrated using a 10 kDa centrifugal concentrator (Sartorius) to 1 mL and loaded onto a Superdex 75 10/300 GL column (GE Healthcare) containing SEC buffer (10 mM MOPS, 5 mM DTT, 1 mM EDTA, pH 7.5) and washed at 0.8 mL·min-1 until protein was eluted. Purity and molar mass were assessed by Coomassie staining and imaging with a Bio-Rad ChemiDoc MP Imaging System following SDS-PAGE (Bio-Rad mini Protean TGX 4–20%). The molar mass of PpEG16.9 was also assessed by intact protein mass spectrometry on a Waters Xevo UPLC-MS system, essentially as previously described (Sundqvist et al., 2007). Proteins were concentrated to approximately 1 mL and stored in SEC buffer at -70°C after snap freezing in liquid nitrogen. 3.2.4 Crystallography trials Sparse matrix crystal screening of PpEG16.9 was carried out in 96-well sitting drop plates using various commercially available screens: Classics Suite, Classics II Suite, JCSG+ Suite, Protein Complex Suite, pH Clear Suite, pH Clear II Suite, and PACT Suite (all from Qiagen). Trays were set up using the Phoenix robot (Art Robbin Instruments) and stored at room temperature or 4°C. Recombinant PpEG16.9 with and without His-tag cleavage by TEV protease were both screened at multiple protein concentrations. Various surface entropy reduction mutants targeting loops connecting various β-strands were also produced by site-directed mutagenesis (Table B.2) using pMCSG53-PpEG16.9 as a template (QuickChange, Agilent Technologies, Santa Barbara, CA, USA), in attempts to promote crystal formation. In particular, we targeted a homologous loop in 76 PpEG16, in which the deletion of a single valine residue enabled crystallization of VvEG16 (McGregor et al., 2017). Unfortunately, extensive crystal screening of all recombinant proteins failed to produce diffraction-quality crystals. 3.2.5 Enzyme kinetics A copper-bicinchoninic acid (BCA) reducing sugar assay was employed to assess the hydrolytic activity of the PpEG16 variants on polysaccharides, essentially as previously described (Arnal et al., 2017). Measurements of temperature and pH optima were performed in 10 min assays containing 1 mg·mL-1 MLG, 16.9 nM PpEG16.9, 0.1 mg·mL-1 BSA, 1 mM NaCl, and 1 mM EDTA. To determine the temperature optimum, 13 reactions were carried out using 50 mM sodium phosphate buffer, pH 7, across the range 9.1-54.0°C. The pH optimum was determined at 37°C with 50 mM citrate buffer 4-6 pH, 50 mM sodium phosphate buffer 6-8.5 pH, or 50 mM glycine buffer 8.5-10 pH, adjusted with NaOH. Initial activity screening was performed using 1.13 mg·mL-1 of individual polysaccharides in a 10-minute assay at 37°C. Michaelis-Menten kinetics were determined using 0.1-5.0 mg·mL-1 polysaccharide in 50 mM sodium phosphate buffer pH 7, 0.01 mg·mL-1 BSA, 0.1 mM NaCl, 0.1 mM EDTA at 37°C for 10 minutes. 16.9 nM of each enzyme was used in substrate specificity, and activity reactions with MLG, 67.5 nM and 33.8 nM were used in the HEC/CMC and xyloglucan, respectively. Non-linear curve-fitting of the Michaelis-Menten equation to the data was performed using OriginPro 2015. Xyloglucan AZCL and hydroxyethyl cellulose AZCL assays (Park and Cosgrove, 2012) were used to verify and normalize endo-(xylo)glucanase activities prior to biomechanical assays (vide infra). 50 μg PpEG16, VvEG16, and PtEG16, were assessed with 1 mg of AZCL 77 polysaccharide in 250 µl of 20 mM sodium acetate, pH 5.5, or 20 mM HEPES, pH 7.5. Samples were incubated for 1 hour at 27o C, with shaking at 1000 RPM. Samples were spun down and filtered before measuring absorbance at 590 nm (Nanodrop 2000, Thermo Scientific). In assays comparing VvEG16 and commercial E-XEGP, the enzymes were incubated independently with 2 mg of AZCL polysaccharide in 1 mL of 20 mM sodium acetate, pH 5.5, for 30 minutes at 27 o C, with shaking at 1000 RPM. Protein concentrations were adjusted to obtain identical A590 readings (Evolution 220 UV-Visible Spectrophotometer, Thermo Scientific) in each assay (1 μg/mL of E-XEGP or 15 μg/mL VvEG16 with AZCL-HEC; 1 μg/mL of E-XEGP or 72.5 μg/mL VvEG16 with AZCL-xyloglucan). 3.2.6 Enzyme product analysis Limit-digest reactions were set up with 100 µg of enzyme and 0.5 mg·mL-1 substrate at optimal conditions of pH 7 and 37°C in 50 mM sodium phosphate buffer for 16 hours (MALDI-TOF MS analysis) and three days (HPAEC-PAD analysis). Each reaction was terminated with 50 mM NaOH and diluted 1:100 with milli-Q H2O before analysis by HPAEC-PAD on a Dionex ICS-5000 system controlled by Chromeleon 7 software. 10 µL injections of samples and standards were separated on a 3×50mm Dionex CarboPac PA200 guard column in series with a 3×250 mm Dionex CarboPac PA200 analytical column. Gradients B and C, previously described by (Larsbrink et al., 2011), were used for the analysis of the MLG and xyloglucan limit-digests, respectively. For MALDI-TOF analysis, samples were mixed with 20 mg·mL-1 2,5-dihydroxybenzoic acid solution at 1:1 ratio applied to Bruker MTP 384 ground steel MALDI plate, and air-dried at room temperature. Samples were analyzed with a Bruker Daltonics Autoflex mass spectrometer. 78 3.2.7 Biomechanical assays Cell wall extension assays were performed as previously described (Cosgrove et al., 2017), using either 4 day-old etiolated chia hypocotyls or 3 day-old etiolated cucumber hypocotyls, abraded with carborundum (Fisher Chemical C-192-500), and heat-inactivated by boiling in deionized water for 15 seconds. Briefly, tissues were clamped into a constant force extensometer at 6 g force (chia) or 20 g (cucumber) in 20 mM sodium acetate buffer (pH 5.5). Hypocotyls were incubated in either 20 mM sodium hydroxide for 20 minutes (chia) or 50 mM CDTA for 30 minutes (cucumber) to sensitize the tissues, which were subsequently washed ten times with deionized water. Samples were then incubated in buffer and allowed to stabilize before the addition of enzymes (protein concentrations are indicated in the corresponding figure legends). Tissue lengths were recorded at 30-second intervals before and after the addition of enzyme. Stress-strain assays were performed on 3-day-old cucumber hypocotyls, abraded with carborundum and heat-inactivated by boiling in deionized water for 15 seconds. Hypocotyls were placed in 1 mL of 20 mM sodium acetate pH 5.5, containing enzyme (protein concentrations indicated in the corresponding figure legends) and incubated for 16 hours at 26°C, shaking at 350 RPM. Samples (5-mm between clamps) were stretched at a constant strain rate of 3 mm/min to a final load of 30 g. 3.3 Results 3.3.1 PpEG16 is representative of non-vascular EG16s EG16 members exhibit high sequence conservation across plant species (Figure 3.1A; see also (Behar et al., 2018)). To juxtapose the previously characterized examples from angiosperms, namely PtEG16 and VvEG16, we selected the ortholog from the model moss P. patens EG16 (PpEG16; previously annotated as PpXTH32 (Yokoyama et al., 2010) for analysis. Indeed, the 79 whole genome-seuqnce of P. patens was the most evolutionarily distant example vis-à-vis angiosperms, available when we initiated our present study (Rensing et al., 2008, Behar et al., 2018). Mosses, including P. patens, diverged from hornworts and liverworts ca. 409 mya in the Devonian era, and these taxa collectively separated from the group that gave rise to current day non-vascular plants 485.4 mya at the beginning of the Ordovician era (Figure 3.1B). Among the mosses, P. patens diverged in the Permian period, ca. 280 mya. In contrast, both V. vinifera and P. trichocarpa diverged in the Cretaceous, 110 and 80 mya, respectively (Morris et al., 2018). Thus, despite a divergence of ca. 500 mya since the split between bryophytes and other land plants, EG16 sequences remain highly conserved, with pairwise amino acid sequence similarities >70% and identities >50% (Figure 3.1A, Figure 3.2A). However, the extent of functional conservation across this evolutionary time and space was unknown. The gene encoding PpEG16 is predicted to have three exons separated by two introns. Detailed analysis of the available nucleotide sequences on Phytozome corresponding to PpEG16 revealed 11 transcripts (Figure B.1) that encode three different predicted protein products (Figure 3.2A). These are designated PpEG16.1, PpEG16.9, and PpEG16.10 after their corresponding transcript number. PpEG16.9 results from the exclusion of the first exon and is the most similar to EG16s from other species (Figure 3.2A). The PpEG16.1 transcript includes all three exons, resulting in a predicted protein with an extended N-terminus. The PpEG16.10 transcript has an exclusion of the first exon, a partial exclusion of the second exon, a gene start in frame +2 of the second exon, and the inclusion of the second intron (Figure B.1). As a result, PpEG16.10 is predicted to have a two amino acid residue insertion at the end of strand 3 and a six amino acid residue insertion in strand 5 versus the tertiary structure of VvEG16 (PDB IDs 5DZE, 5DZF, 5DZG, 5SV8; (McGregor et al., 2017)) (Figure 3.2A). Otherwise, the three predicted PpEG16 80 variants have identical sequences in the C-terminal 153 amino acid residues of the proteins (Figure 3.2A; starting at G55), including the β-strand presenting the catalytic residues (EIDFE, underlined, E88-E92). The minimal variant PpEG16.9 aligns with PtEG16 and VvEG16 with pairwise sequence similarities of 68% and 69%, respectively (Figure 3.1A), allowing the production of a structural homology model based on the crystal structure of VvEG16 (Figure 3.2B, Figure B.2). Figure 3.2. PpEG16 sequence and structure analysis. (A). Structure-based alignment of PpEG16 and other characterized EG16s (also available as Supplemental Material 3.4). Active-site amino acid residues are highlighted in purple. (B). Wall-eyed (divergent) stereo view of a superposition of the active-sites of VvEG16 (PDB ID 5DZG) and a structural homology model of PpEG16 (Supplemental File 3.2). See Figure B.2 for a global superposition. 81 Synthetic cDNA sequences corresponding to N-terminal 6xHis-tagged forms of the three PpEG16 variants were expressed in E. coli, resulting in the production of soluble proteins. Following purification by nickel affinity chromatography and size exclusion chromatography (Figure B.3), yields of 18.1 mg, 15.4 mg, and 5 mg per litre of culture medium were obtained for PpEG16.1, PpEG16.9, and PpEG16.10, respectively. After purification, all three proteins appeared as major bands whose migrations on SDS-PAGE were consistent with the expected molar masses; some evidence of protein degradation was also observed (Figure B.3A). PpEG16.9 was detected as a single peak by intact protein mass spectrometry (Figure B.3B). 3.3.2 PpEG16 is predominantly a mixed-linkage -glucanase The substrate specificity of the three PpEG16 variants was initially screened on a series of -glucan, -xylan, and -mannan polysaccharides as representative potential substrates (Figure B.4). The insertion variant PpEG16.10 was poorly active or inactive on all substrates, precluding conclusions on substrate specificity. In contrast, variants PpEG16.1 and PpEG16.9 were efficient endo-glucanases, with the highest specific activities on barley MLG (v0·[E]t-1 values of 289.5 min-1 and 373.3 min-1, respectively) (Figure B.4). Both variants were also active on the soluble artificial cellulose ((1,4)-glucan) derivatives carboxymethyl cellulose (CMC) and hydroxyethyl cellulose (HEC), yet with less than 20% of the specific activity on MLG (Figure B.4). Notably, the relative specific activity on tamarind galactoxyloglucan, which comprises a (1,4)-glucan backbone with extensive (1,6)-xylosyl branches, was much lower than for the angiosperm PtEG16 (Eklöf et al., 2013) and VvEG16 (McGregor et al., 2017). For example, the relative specific activity of PpEG16.9 on 1.13 mg/mL xyloglucan was 3% of the value on MLG (Figure B.4), which can be compared with a ratio of kcat values of 1:2 (MLG to xyloglucan) for VvEG16 82 (McGregor et al., 2017). The variants PpEG16.1 and PpEG16.9 had marginal but detectable activity on konjac glucomannan (kGM) and wheat arabinoxylan (wAX). However, the relative activities of PpEG16.9 on these polysaccharides were only 0.84% and 0.35% that of MLG, respectively (Figure B.4). Given the similar activities of PpEG16.1 and PpEG16.9 and the greater similarity of the length of the PpEG16.9 polypeptide with other EG16 members (Figure 3.2), PpEG16.9 was selected for further detailed kinetic characterization. Thus, temperature- and pH-rate profile measurements indicated optimal in vitro assay conditions of pH 7 and 37°C in 50 mM sodium phosphate buffer, using MLG as a substrate (Figure B.5). The preceding activity measurements were performed with EDTA in the assay buffer, based on preliminary observations. Indeed, PpEG16.9 activity was significantly lower (ca. 5-fold) in the absence of 0.1 mM EDTA in the assay (Figure B.6). EDTA also had a slight positive effect on the stability of PpEG16.9 at 4°C: Without EDTA, activity is reduced by 50% over 2 hours, while with EDTA, the activity is only reduced by 10% over the same period. Therefore, all kinetic assays were carried out with a final concentration of 0.1 mM EDTA. This phenomenon has been previously observed for other endoglucanases, in which it has been attributed to enhanced stabilization of the protein structure, although the mechanism is unknown (Naika and Tiku, 2011). Michaelis-Menten kinetics were determined for each of the four substrates for which significant specific activity was observed (Figure 3.3). Concordant with our initial observations, the highest kcat value (703 min-1) and lowest KM value (1.04 mg·mL−1) were observed with MLG, yielding an apparent kcat/KM value of 675 mL·mg−1 s−1 (Table 3.1). The artificial substrates HEC and CMC had apparent kcat/KM values 6-fold and 10-fold lower than that of MLG. Most strikingly, the kcat/KM value for xyloglucan was 70-fold lower than that of MLG. This latter ratio contrasts 83 that of VvEG16 (McGregor et al., 2017), which is only 13-fold less specific for xyloglucan (Table 3.1). Figure 3.3. Michaelis-Menten kinetics of PpEG16.9.The release of reducing ends from barley mixed-linkage β-glucan (A), hydroxyethyl cellulose (B), carboxymethyl cellulose (C), and tamarind xyloglucan (D) was quantified by a copper-bicinchoninic acid end-point assay. Each point is an average value of triplicate measurements. Error bars represent the standard deviation. Lines represent the non-linear fitting of the Michaelis-Menten equation to the data. The limit-digest products of PpEG16.9 acting on MLG and xyloglucan were determined by HPLC and MS (Figure 3.4). By comparison with standards, four main products were identified by HPAEC-PAD in an extended (3 day) digest of barley MLG: glucose (G), cellobiose (GG), cellotriose (GGG), and the mixed-linkage tetrasaccharide G3GGG [β-D-Glcp-(1,3)-β-D-Glcp-(1,4)-β-D-Glcp-(1,4)-β-D-Glcp] (Figure 3.4, cf. Figure B.7). These results were supported by 84 MALDI-TOF MS analysis, which clearly indicated the presence of hexosyl di-, tri-, and tetrasaccharides, as well as a Hex7 species likely resulting from incomplete enzymatic hydrolysis (Figure 3.4). This product distribution, especially the observation of the G3GGG tetrasaccharide bearing a (1,3) linkage at the non-reducing end, mirrors those observed for VvEG16 and PtEG16 (Eklöf et al., 2013, McGregor et al., 2017). Table 3.1. PpEG16 and VvEG16 apparent Michaelis-Menten constants for polysaccharide hydrolysis Substrate Enzyme kcat, app (min−1) KM, app (mg·mL−1) kcat/KM, app (mL·mg−1 min−1) MLG PpEG16.9 703 ± 24 1.04 ± 0.12 675 VvEG16* 163 ± 5 0.36 ± 0.02 452 HEC PpEG16.9 211 ± 10 1.88 ± 0.27 112 VvEG16(ΔV152)* 10.4 ± 0.8 1.1 ± 0.2 9.45 CMC PpEG16.9 203 ± 11 3.15 ± 0.43 64.4 VvEG16(ΔV152)* 17 ± 2.2 2.8 ± 0.5 6.07 XyG PpEG16.9 53.2 ± 6.5 5.37 ± 1.2 9.91 VvEG16* 70 ± 6 2.1 ± 0.4 33.3 *Values from (McGregor et al., 2017). Error values represent those obtained from non-linear fitting of the Michaelis-Menten equation to the data. Incubation of PpEG16.9 with tamarind xyloglucan produced the canonical oligosaccharide products resulting from hydrolysis at unbranched glucosyl residues in the backbone (Eklöf et al., 2012), i.e., XXXG, XLXG, XXLG, and XLLG, as well as small amounts of larger combinations of these oligosaccharides due to incomplete digestion (Figure 3.4, cf. Figure B.7). Although some minor signals corresponding to smaller oligosaccharides were observed by MS (16 hours, Figure 3.4D) and HPLC (3 days, Figure B.7) after extended incubation, there is little evidence of hydrolysis at branched xyloglucan backbone residues, which has been observed for a limited number of microbial endo-xyloglucanases (Arnal et al., 2019). As for MLG, this cleavage pattern recapitulates those observed for VvEG16 (McGregor et al., 2017) and PtEG16 (Eklöf et al., 2013). 85 Figure 3.4. Product analysis of PpEG16.9. 100 µg of PpEG16 enzyme was incubated with 0.5 mg·mL-1 mixed-linkage β-glucan (A, B) or xyloglucan (C, D) to produce limit digest products, assessed by HPAEC-PAD (A, C) and MALDI-TOF (B, D). 3.3.3 EG16 orthologs weaken plant cell walls ex vivo To gain insight into the potential physiological function of PpEG16, we applied biomechanical assays that were previously established to measure the activity of expansins (McQueen-Mason and Cosgrove, 1994, McQueen-Mason and Cosgrove, 1995), XET (McQueen-Mason et al., 1993, Saladié et al., 2006), and microbial endo-(xylo)glucanases (Park and Cosgrove, 86 2012) on plant cell walls ex vivo. Thus, hypocotyl wall extension (creep) under a constant load or mechanical stress under a constant strain rate were measured in an extensometer (Cosgrove et al., 2017). Initial experiments indicated no stimulation of creep by PpEG16 or by the orthologs PtEG16 and VvEG16 when applied at 200 μg/mL to chia or cucumber hypocotyls prepared in the standard manner (i.e., frozen, thawed and heated to inactivate endogenous enzymes and expansins) (data not shown, cf. Figure 3.5E). For comparison, mixed-function microbial endo-(1,4)-glucanases/xyloglucanases, e.g., Paenibacillus sp. E-XEGP or Hypocrea jecorina Cel12, induced sustained creep following an initial lag phase (Park and Cosgrove, 2012). Wall samples were then chemically weakened in further attempts to elicit an effect on wall mechanics. For chia hypocotyls pretreated with 20 mM NaOH, all three enzymes induced extension after a variable lag of ~10-50 min; the extension was not stable but quickly led to mechanical failure (Figure 3.5A-C). Distinctly, PpEG16 had no effect on the comparatively thicker cucumber hypocotyls, which were pretreated with 50 mM CDTA (Figure 3.5E) to sensitize them to these enzymes. In contrast, the orthologs VvEG16 and PtEG16 eventually induced an erratic extension followed by failure (Figure 3.5F-G). Attempts to obtain sustained creep by adjusting the enzyme concentrations or to remove the enzymes upon the start of the extension were unsuccessful: In all cases, extension led to breakage. 87 Figure 3.5. Extension assays under constant load with sensitized hypocotyl walls. EG16 (200 µg/mL) was added at the time point indicated by arrows. Cucumber was sensitized by pretreatment with CDTA, chia with NaOH. At least three biological replicates were used for each enzyme. We next assessed the effects of PpEG16, VvEG16, and PtEG16 on the stress-strain behaviour of cucumber walls following 16-hour treatments with a constant protein loading of 200 88 µg/ml. PpEG16 and PtEG16 had no effect, while VvEG16 increased the plastic compliance by 70% and the elastic compliance by 13% (Figure 3.6). For comparison, treatment of cucumber hypocotyls with 0.1 μg/mL of the fungal HjCel12A for 1 hour increased the plastic compliance by 70% and the elastic compliance by 30% (Park and Cosgrove, 2012). Subsequent experiments in which PpEG16 and PtEG16 enzyme loadings were normalized by activity versus 200 µg/mL VvEG16 (PpEG16, 3 mg/mL; PtEG16, 297 µg/mL) likewise failed to evidence any changes in elastic and plastic compliances by these plant endo-glucanases (Figure B.8-B.10). In contrast, the equivalent activity of 2.76 µg/mL E-XEGP significantly altered both the elastic and plastic compliance of cucumber hypocotyls, consistent with previous results (Park and Cosgrove, 2012). The addition of DTT and EDTA, which enhance the stability and activity of PpEG16 in vitro, did not affect the results of the ex vivo biomechanical assay (Figure B.11). Figure 3.6. Stress-strain analysis of cucumber hypocotyls. 200 µg/mL of each protein was used. Error bars indicate the standard errors (n = 13). The asterisk indicates significance based upon Student’s t-test (two-way, p < 0.05). 89 3.4 Discussion Since the initial identification of the EG16 clade as sister to the XTH gene products in GH16s, the enzymology and structural biology of EG16 members has been restricted to two angiosperm orthologs (Eklöf et al., 2013, McGregor et al., 2017). In the present study, we leveraged our comprehensive census of EG16s in available genomes and transcriptomes (Behar et al., 2018, Jiao et al., 2020) to demonstrate that the biochemical properties of these enzymes are essentially conserved across 500 million years of plant evolution, using the ortholog from the model moss P. patens as a non-angiosperm example (Figure 3.1). Notably, PpEG16 is highly similar to, and thus representative of, other moss orthologs (Figure 3.1) (Behar et al., 2018). The data shows that the substrate specificity of the moss EG16 is essentially conserved vis-à-vis the orthologs from poplar and grape, but with some notable differences. Although the requirement of a protein reducing agent to maintain the stability of PtEG16 precluded quantitative reducing-sugar activity assays, the poplar enzyme exhibited broad specificity towards both the linear MLG and the highly branched xyloglucan at qualitatively similar capacities (Eklöf et al., 2013). The grape ortholog VvEG16, on the other hand, was kinetically tractable, with a preference for MLG over xyloglucan evidenced by an order of magnitude difference in kcat/KM values for these cell wall polysaccharides (Error! Reference source not found.) (McGregor et al., 2017). The apparent kcat/KM values for MLG are comparable between the moss and the grape orthologs, yet PpEG16 exhibited a four-fold lower kcat/KM value for xyloglucan (Error! Reference source not found.). Thus, the 70-fold difference in kcat/KM values of PpEG16 for MLG over xyloglucan indicates an even stronger bias against the highly branched -glucan. The biological significance of this is unclear, as P. patens is not known to contain large amounts of MLG (Roberts et al., 2012). One possibility is that the native substrate for PpEG16 is amorphous cellulose, for which 90 MLG is a natural proxy in our in vitro assays. Notably, PpEG16 exhibited a higher relative activity than VvEG16 on the artificial amorphous cellulose derivatives HEC and CMC. It is also possible that the apparent lower activity of PpEG16 on xyloglucan might be due to the use of the readily available tamarind (eudicot) XXXG-type xyloglucan in the assays, whereas native moss xyloglucan is of the less-branched XXGG-type (Schultink et al., 2014). A further possibility is that arabinoglucan, a polysaccharide recently discovered in P. patens (Roberts et al., 2018), could be a substrate for PpEG16. Although the hydrolysis of eudicot xyloglucan by PpEG16 produced the well-known series of Glc4-based oligosaccharides through cleavage at the unbranched glucosyl residues, the hydrolysis products of MLG deserve special comment. Like VvEG16 and PtEG16, PpEG16 produced the distinct tetrasaccharide G3GGG (Figure 3.4 and Figure B.7). This product results from the site-specific cleavage of a β(1,4) bond to a β(1,3)-linked Glc residue in MLG (e.g., G3GGG↓G3GGG). Thus, this mode involves specific recognition of a β(1,3)-linkage spanning enzyme subsites +1 and +2 (subsite nomenclature according to (Davies et al., 1997)). In contrast, canonical bacterial MLGases (licheninases) require that the β(1,3)-linkage span subsites -2 and -1, resulting in the production of the oligosaccharides GG3G and GGG3G (Planas, 2000, Tamura et al., 2017, Déjean et al., 2020). The observation of glucose, cellobiose, and cellotriose, and no other mixed-linkage oligosaccharides, suggests hydrolysis of both β(1,3) bonds (analogous to laminarinases (Badur et al., 2020, Déjean et al., 2020)) and β(1,4) bonds by PpEG16, as was observed for PtEG16 (Eklöf et al., 2013) and VvEG16 (McGregor et al., 2017). Despite extensive screening, we were unable to produce diffraction-quality crystals of the PpEG16 variants alone or in complex with oligosaccharides. Hence, we generated a structural homology model (Figure 3.3 and Figure B.7) based on the experimentally determined structure 91 of VvEG16 (McGregor et al., 2017). As expected, this model had the characteristic -jelly-roll architecture, with gross differences restricted to loops distant from the active-site cleft (Figure B.2). The catalytic residues and residues lining the active-site cleft were also found to be conserved and positioned virtually identically in PpEG16 vis-à-vis VvEG16, with the exceptions of Asp47 in place of Gly43, Ser79 in place of Asn75, and Trp180 in place of Tyr175 (Figure 3.2B). Yet, none of these differences present an obvious steric barrier, in the absence of an experimental three-dimensional structure, that would explain the lower relative activity of PpEG16 towards xyloglucan. Previous studies found that microbial endo-(1,4)-glucanases with the ability to hydrolyze both xyloglucan and soluble cellulose derivatives induced creep (slow, sustained extension) of cucumber hypocotyl and onion epidermal walls, whereas enzymes selective for either substrate lacked this ex vivo activity (Zhang et al., 2019). Despite possessing a similar mixed endo-glucanase/xyloglucanase activity, PpEG16, as well as the orthologs from poplar and grape, did not induce creep but rather caused the abrupt, catastrophic failure that was dependent upon hypocotyl source and pretreatment. Also distinct from the microbial enzymes, the EG16 orthologs produced little to no softening, i.e., increases in elastic or plastic compliances, in these cell walls, even when applied at high protein loadings to normalize enzyme activities. As the biomechanical actions of the EG16 enzymes are both markedly smaller and distinct, we conclude that they operate on the cell wall matrix through a different mechanism than that proposed for their microbial counterparts (Park and Cosgrove, 2012). In this regard, the EG16 orthologs are also distinguished from the plant expansins and their bacterial homologs, which stimulate wall creep (McQueen-Mason and Cosgrove, 1994, McQueen-Mason and Cosgrove, 1995, Cosgrove, 2017). 92 In this context, it is worthwhile to compare the ex vivo effects of PpEG16, VvEG16, and PtEG16 with the limited data available on plant XTH gene products, which has been long suggested to catalyze plant cell wall expansion (Rose et al., 2002). Despite this association, exogenous addition of xyloglucan endo-transglycosylases (XET) had no detectable effect on the mechanical properties of cucumber hypocotyls ex vivo, viz. they did not cause wall extension nor softening, including after the addition of exogenous oligosaccharides as alternate acceptor substrates (McQueen-Mason et al., 1993, Saladié et al., 2006). Here, we observed analogous negative results for PpEG16 on the same substrate (Figure 3.5, Figure 3.6, and Figure B.10), whereas this enzyme caused breakage of the much smaller chia hypocotyls (Figure 3.5A). On the other hand, VvEG16 and PtEG16 caused the breakage of both cucumber and chia hypocotyls (Figure 3.5). One explanation for these different results could be the lower activity of PpEG16 on xyloglucan, which is the predominant matrix polysaccharide in these eudicot tissues (Carpita and McCann, 2000, Scheller and Ulvskov, 2010). Regardless, our observations of catastrophic tissue failure in the absence of detectable softening are confounding and indicate that the ultrastructural effects of these endogenous plant enzymes remain opaque. In this context, it is notable that the genetic deletion of two paralogous XEH in Arabidopsis thaliana (Group III-A enzymes encoded by AtXTH31 and AtXTH32) resulted in the elimination of detectable endo-xyloglucanase activity in vivo, failed to cause growth or developmental phenotypes (Kaewthai et al., 2013). In conclusion, the biochemical analysis presented here provides new insight into the range of specificities exhibited by plant EG16 orthologs spanning diverse evolutionary lineages. Although the physiological functions of this unique group of enzymes remain to be revealed in vivo, these results will inform future genomic and biological studies across a wide range of plant taxa. 93 Chapter 4: Physcomitrella patens endo-glucanase 16 is involved in the cell wall development of young tissue 4.1 Introduction The cell wall is an essential and defining feature of all plant cells, giving the cell various shapes and contributing to structure, strength, and functionality, while also providing a defence against pathogens (McNeil et al., 1984, Cosgrove, 2005, Wolf et al., 2012). The primary cell wall comprises the polysaccharide components cellulose, hemicelluloses such as xyloglucan, and pectins, which form a complex network that requires maintenance and modification during growth and response to the environment (Gigli-Bisceglia et al., 2020). Diverse proteins and enzymes contribute to cell wall morphogenesis (Jamet and Dunand, 2020), prominent of which are the xyloglucan endo-transglycosylase/hydrolase (XTH) gene products. These enzymes are a subfamily of glycoside hydrolase family 16 (GH16) (Viborg et al., 2019), and individual members catalyze the rearrangement (xyloglucan endotransglycosylases, XETs, EC 2.4.1.207) or irreversible hydrolysis (xyloglucan endohydrolases, XEHs, EC 3.2.1.151) of xyloglucan chains (Baumann et al., 2007). Within this subfamily, enzymes that catalyze polysaccharide hetero-transglycosylation have also been discovered, which results in the production of xyloglucan-pectin hetero-polysaccharides (Stratilová et al., 2020a). Thus, XTH gene products, which comprise groups of 20-60 homologs in land plants, have been implicated in wall loosening, wall strengthening, and integration of nascent xyloglucan into the wall (Rose et al., 2002, Miedes and Lorences, 2009, Eklöf and Brumer, 2010, Cosgrove, 2016). A distinct clade of plant GH16 endo-glucanases (EG16), which is sister to the XTH gene products, was revealed by detailed bioinformatic and biochemical analysis (Eklöf et al., 2013). EG16 members lack the characteristic C-terminal extension of XTH gene products (PFAM 94 XET_C, PF06955) and exhibit distinct activity, hydrolyzing both mixed-linkage β-glucan (MLG) and xyloglucan (Eklöf et al., 2013). Notably, this activity is conserved in distantly related members of black cottonwood (Populus trichocarpa) (Eklöf et al., 2013), grapevine (Vitis vinifera) (McGregor et al., 2017), and spreading earthmoss (Physcomitrella patens) (Chapter 3). Moreover, an extensive cross-genome/transcriptome analysis revealed that EG16 genes are generally present in members from all plant clades, but are missing from many eudicot genomes, particularly asterids and rosids such as Arabidopsis (Behar et al., 2018). Further analysis of green algal genomes revealed that EG16 genes were present in the most phylogenetically distant green algae species to land plants, while XTH genes were absent (Jiao et al., 2020), supporting the proposal that EG16 and XTH members share a common ancestor that is EG16-like (Eklöf et al., 2013). A particular curiosity is that EG16 members are present as single orthologs in all land plants surveyed thus far, in striking contrast to multi-gene XTH families, which may suggest a specialized role for EG16 gene products. Despite extensive biochemical and structural characterization (Eklöf et al., 2013, McGregor et al., 2017) (Chapter 3), the biological function of EG16 orthologs is presently unknown. Since EG16 genes are strongly represented in non-angiosperm genomes (Behar et al., 2018), we selected the moss P. patens as a well-established model organism (Cove, 2005, Quatrano et al., 2007, Rensing et al., 2020) to explore the role of an exemplar EG16 in vivo. The gene encoding P. patens EG16 (PpEG16) was initially identified as an XTH (PpXTH32) by sequence similarity in an early genomic survey (Yokoyama et al., 2010). Subsequently, refined phylogenetic analysis has shown that PpEG16 and all EG16 orthologs comprise a sister clade to the XTH gene products (Eklöf et al., 2013, McGregor et al., 2017, Behar et al., 2018). Notably, the PpEG16 sequence lacks the characteristic C-terminal extension of the 95 XTHs (Johansson et al., 2004, Baumann et al., 2007, Eklöf et al., 2013, McGregor et al., 2017) and comprises the common EG16 motifs (Behar et al., 2018). Recently, enzymology of PpEG16 demonstrated activity analogous to previously characterized EG16s, i.e., predominant hydrolytic activity on MLG and secondary activity on xyloglucan and soluble cellulosic substrates (Chapter 3). Although P. patens cell walls, like those of other mosses and most land plants, contain the matrix polysaccharide xyloglucan, MLG is absent (Popper et al., 2011). As MLG is limited to a few plant families (e.g., graminaceous monocots), the MLGase activity of EG16s is enigmatic. The goal of this study is to gain insight into the biological role of PpEG16. We performed diverse techniques to investigate this, including expression analyses, immunolocalization, and examination of loss-of-function growth phenotypes. The research revealed that PpEG16 is expressed in young, developing protonema of P. patens, and EG16s localizes to the cell wall of growing tips of protonema, where there is a high concentration of xyloglucan. Using a reverse genetics approach, we also demonstrate that deletion of PpEG16 causes an increase in growth and premature senescence. 4.2 Materials and methods All materials and chemicals were purchased from Sigma-Aldrich unless otherwise stated. Protocols were adapted from the Bezanilla Lab Moss Methods Manual, available at URL https://sites.dartmouth.edu/bezanillalab/files/2018/01/Moss-Manual.pdf (Rensing et al., 2020). 4.2.1 Data sources JGI Phytozome was used for accessing the Physcomitrella genomic sequence (accession number Pp3c21_9420V3.9) (Goodstein et al., 2012). Expression data for PpEG16 was obtained from the P. patens eFP browser (accession number Pp1s353_23V6.1) (Winter et al., 2007, Ortiz-Ramirez et al., 2016). 96 4.2.2 Growth conditions P. patens (ecotype Gransden) protonemata were grown at 25°C under continuous light (73.6 µmol·s-1) on plates containing minimal media supplemented with ammonium tartrate (“PpNH4 media”: 3.39 mM Ca(NO3)2·4H2O, 1.03 mM MgSO4·7H2O, 1.84 mM KH2PO4, 82.29 µM FeSO4·7H2O, 2.72 mM diammonium tartrate, 0.7% agar (w/v) and micro-elements mix (10 µM H3BO3, 220.27 nM CuSO4·5H2O, 1.97 µM MnCl2·4H2O, 231.16 nM CoCl2·6H2O, 191.24 nM ZnSO4·7H2O, 1.69 nM KI, 103.33 nM Na2MoO4·2H2O) covered in cellophane (1080; Research Products International) and sealed with micropore tape (1530-1; 3M). Tissue culture plants were maintained by a weekly collection of tissue into round bottom plastic tubes containing sterile dH2O, homogenization using a bead beater (BioSpec Mini-Beadbeater) with a single 3 mm glass bead, and growth on fresh PpNH4 plates. 4.2.3 Plasmid construction and moss transformation The P. patens genome was extracted by grinding ca. 100 mg of moss tissue with a blue polypropylene pestle (Sigma Aldrich) in 1.7 mL tube with 400 µL extraction buffer (0.2 M Tris-HCL, pH 9, 0.4 LiCl, 25 mM EDTA, 1% SDS). After centrifugation at 20,000 x g for 5 minutes, the supernatant was gently mixed with an equal volume of isopropanol. The mixture was centrifuged for an additional 10 minutes before the supernatant was discarded. The pellet was left to dry for 20 minutes at ambient temperature and finally resuspended with 400 µL TE buffer (10 mM Tris pH 8, 1 mM EDTA). Homologous recombination was used for gene targeting in P. patens. For PpEG16 knockout mutants, PCR was performed on 2 µL of P. patens genomic DNA to clone two amplicons containing DNA from the 5' and 3' ends of the EG16 gene (Table C.1). The 5' flanking region of the PpEG16 gene was inserted into the pBKS plasmid containing a Geneticin-resistance cassette 97 using restriction enzymes NotI and BamHI, while the 3' flanking region was inserted using SalI and KpnI (New England Biolabs). Plasmids for the complementation of EG16 (in a background of ΔPpEG16-1 (EG16 knockout line)) were constructed to insert PpEG16 with a C-terminal mCherry under the native PpEG16 promoter. The 3' region of the PpEG16 gene was inserted into the pMDC111-mCherry Gateway cloning vector with SacII and PsiI restriction enzymes. The 5' region of the PpEG16 gene was amplified by PCR with the full-length PpEG16 gene and inserted into the Gateway Donor vector pDONR 207 by a BP reaction (ThermoFisher Scientific) before utilizing an LR reaction to transfer the amplicon into the pMDC111-mCherry already containing the 3' region. PCR reactions were run in Bio-Rad C1000 Thermal Cycler. Plasmids were transformed into competent E. coli DH5α cells via heat shock at each cloning step, and then grown on LB media supplemented with appropriate selection reagent. Sequence accuracy was confirmed by Sanger sequencing. Preparation of DNA for polyethylene-glycol (PEG)-mediated transformation of P. patens and selection of transformants was carried out as previously described (Wu et al., 2011, Wu and Bezanilla, 2014). Selection plates contained 25 µg/mL geneticin for PpEG16 deletion lines and 30 µg/mL hygromycin for PpEG16-mCherry lines. 4.2.4 Genotyping Genomic DNA was isolated as described above, and 2 µL of the extraction was used as the PCR template. The plasmids used for transformations served as templates in positive controls. The resulting PCR products were loaded into 1% agarose gel (w/v) supplemented with 0.002% RedSafe (v/v) (FroggaBio) and run at 80 V for 50 minutes alongside 1kb DNA ladder (FroggaBio) (Figure C.2). The PCR products were sequenced by Sanger sequencing to confirm accuracy. 98 4.2.5 Growth assays Growth assays were carried out by placing ca. 1 mm pieces of 7-day-old protonema on plates containing minimal media ("PpNO3 media": 3.39 mM Ca(NO3)2·4H2O, 1.03 mM MgSO4·7H2O, 82.29 µM FeSO4·7H2O, 1.84 mM KH2PO4 pH 7, micro-elements mix, 0.7% agar (w/v)) and monitoring growth at weekly intervals. Dark growth assays were similarly prepared, but the tissues were placed into square Petri dishes containing PpNO3 media supplemented with 5 mM diammonium tartrate and 2% sucrose (w/w), grown in continuous light for ten days, and then the plates were flipped vertically, and completely concealed from light for either two or four weeks. The assays were repeated three times. Plates were photographed using a Bio-Rad ChemiDoc MP Imaging System, while ImageJ was utilized in image analysis (Schneider et al., 2012), and Microsoft Office Excel 2019 and OriginPro 2015 were used for analysis and graphing. 4.2.6 Sample preparation and microscopy procedures For live-imaging microscopy, 4-week old gametophores and 7-day old protonema tissue of wild-type and PpEG16-mCherry transgenics were collected into 8% mannitol (w/v) solution or dH2O. For immunofluorescence microscopy, 7-day-old protonema tissue was incubated with fixative (50 mM PIPES pH 7.5, 5 mM MgSO4, 5 mM EGTA pH 7.5, 3.7% paraformaldehyde) for 35 minutes with gentle agitation and subsequently washed twice with TBST buffer (20 mM Tris pH 7.6, 0.5 M NaCl, 0.2% Tween-20 (w/v)). Tissues were then ethanol-bleached with gradually increasing ethanol concentrations of 25% (v/v), 50%, and 100% for 10 minutes each under gentle agitation, then gradually decreasing concentrations of the same solutions. Finally, the tissues were washed with TBST. Next, the tissues were separated into different tubes and then incubated with 5% skim milk powder (w/v) in TBST buffer for 1 hour at room temperature under gentle agitation. The tubes were centrifuged at 5,000 x g for 3 minutes, and the supernatant was discarded and 99 replaced with 54 µL of one of the following primary antibodies diluted 1:36 in TBST: anti-mCherry antibody (PA5-34974, ThermoFisher Scientific); anti-xyloglucan (LM15, ab190146; Abcam); anti-α-tubulin (T5168, Millipore Sigma); anti-(1,3;1,4)-β-glucan (BG1/BS 400‐3, Biosupplies Australia); or TBST only. The samples were then left at 4°C under gentle agitation overnight. The samples were then centrifuged, and the pellet was washed in TBST for 10 minutes twice before one of the next secondary antibodies was added at 1:100 dilution in TBST: Alexa Fluor 488 anti-rat (A-11006), anti-mouse (A-11001), or anti-rabbit (A-11008) from Invitrogen. The mixture was incubated for 1.5 hours at room temperature with slight agitation. Finally, the tissues were centrifuged again and washed twice with TBST for 10 minutes each. For cell wall staining, wild-type protonema tissue was incubated with 0.1% calcofluor-white (fluorescent brightener) (w/v) in 8% mannitol (w/v) for 1 hour, then washed with 8% mannitol for 30 minutes. Tissues were mounted on microscope slides (FroggaBio) in TBST (immunofluorescence samples), 8% mannitol (w/v), or dH2O (live-tissue samples) and covered with a coverslip. Data collection was accomplished using 10x/NA 0.4/Air, 40x/NA 0.95/Air, and 60x/NA 1.1; WD 1.5mm lenses on an Olympus FV1000 Laser Scanning microscope. 559 nm laser was used for excitation of the live tissue, and the emission was limited to 590-640 nm to target mCherry fluorescence. For immunofluorescence imaging, 473 nm laser was used for excitation of the Alexa Fluor 488 probe, and default settings were used for emission at 500-600 nm. Calcofluor-white stained tissues were mounted in 8% mannitol and excited with a 405 nm laser with cyan fluorescent protein (CFP) dye setting for emission at 400-500 nm. 100 4.3 Results 4.3.1 PpEG16 is expressed in young tissues To gain initial insight into the potential biological roles of PpEG16, we explored existing transcriptomic data (Ortiz-Ramirez et al., 2016). PpEG16 is most highly expressed in protoplasts and protonema (comprising chloronema and caulonema) (Figure C.1A). Intriguingly, expression in the faster-growing caulonema (Rensing et al., 2020) appears to be higher than in the chloronema (Figure C.1B). Several-fold lower expression levels were observed in the gametophore, rhizoids, and sporophyte S3 stage, with still lower expression in earlier and mature sporophyte developmental stages (Figure C.1B) (Ortiz-Ramirez et al., 2016). Together, these data suggest a role for PpEG16 in cell wall expansion and morphogenesis of young tissue. Also consistent with this transcriptomic data, the early promoter-GUS analysis indicated that PpEG16 (then known as PpXTH32) is strongly expressed in developing protonema, apical tissues of the gametophore, young leaves, and rhizoids (Yokoyama et al., 2010). 4.3.2 Growth phenotyping of ΔPpEG16 transgenics To further probe the physiological role of PpEG16, complete knockout transgenics were made by homologous recombination, producing two independent lines, ΔPpEG16-1 and ΔPpEG16-2. The deletion of PpEG16 was confirmed by genotyping (Figure C.2A). For phenotyping, 7-day-old protonema pieces of the wild-type and knockout lines were grown for five weeks (Figure C.2). During this period, no apparent gross morphological abnormalities of the colonies were observed, e.g., there was no irregular leaf morphology in gametophores, differences in the number of gametophores, or overall plant shape. On the other hand, a slightly enhanced growth phenotype was observed early in the colony growth period. ΔPpEG16-1 transgenic colonies were consistently larger than the wild-type starting from week 1 until the end of the assay. 101 The phenotype was most pronounced in the early stages of growth when the ΔPpEG16-1 plant area ranged from 119% larger at week 1 (p-value = 0.01) to 132% at week 2 (p-value = 2.28·10-8) (Figure 4.1A-B, Figure C.3). An increased growth rate was only consistently observed for the second mutant line, ΔPpEG16-2, at week one. For week 2, ΔPpEG16-2 plants were only statistically larger in two of the three assays (Figure C.4A-B). Similar variability was observed for ΔPpEG16-1 at the later time points, where it was statistically significantly larger than wild-type in two out of three assays (Figure C.4B-C). These data indicate that, within the variability of the colony growth assay, the loss of PpEG16 resulted in larger colonies, especially earlier in the colony growth. 102 Figure 4.1. Growth assay of wild-type, ΔPpEG16, and PpEG16 complementation moss lines. 7-day-old 1 mm pieces of protonema were grown into colonies on minimal media under continuous light at 25°C, and their growth was evaluated by calculating plant area. Boxes represent data points between the 25th and 75th percentile, the whiskers correspond to the minimum, and maximum measurements and lines inside the boxes represent the mean. Significance was evaluated by a 2-tailed Student’s t-test of mutant compared to wild-type. (A). Summary of growth of wild-type and PpEG16 deletion lines (N = 18 for each line). Expanded data is shown in Figure C.4. (B). Representative image of growth assay plate at 2 weeks. Representative growth for each week is presented in Figure C.3. (C). PpEG16-mCherry complementation growth assay including wild-type and four PpEG16-mCherry plants. (D). Growth plate after 2 weeks, showing representative plant sizes. If the larger-sized colony phenotype seen in ΔPpEG16 is due to loss of function of PpEG16, then restoring a wild-type copy of the gene into the mutants is predicted to lead to smaller colonies. The ΔPpEG16-1 line was complemented by inserting PpEG16-mCherry into the genome 103 under the native PpEG16 promoter, producing six lines (Figure C.2B). A growth assay using four of these lines (PpEG16-mCherry 2-6, PpEG16-mCherry 2-7, PpEG16-mCherry 2-10, and PpEG16-mCherry 2-12) indicated similar growth rates to the wild-type in three of the four lines, demonstrating that the wild-type copy of the gene could rescue the mutant phenotype. One line, PpEG16-mCherry 2-12, consistently grew slightly larger than the other lines (p-value = 7.95·10-7 at week 3) (Figure 4.1C-D), suggesting that it retained the mutant phenotype. To test whether PpEG16 is involved in light-independent growth, both the deletion lines and wild-type were grown in continuous light to establish, then transferred to vertical growth conditions in the dark. The two ΔPpEG16 were statistically significantly longer than the wild-type after two weeks, measuring 1.83 ± 0.33 cm (p-value = 0.005) and 1.83 ± 0.24 cm (p-value = 0.01) in comparison to 1.74 ± 0.3 cm of the wild-type (Figure 4.2A). Although the growth phenotype difference was small, it was consistent across three biological replicates, and it disappeared after four weeks of growth (Figure 4.2B), analogous to observations during extended light-grown conditions. Thus, the loss of PpEG16 was consistently associated with increased growth, suggesting a potential role for PpEG16 in restricting growth during moss development. 104 Figure 4.2. Dark growth assay of wild-type and ΔPpEG16 moss lines. 7-day-old 1 mm protonema pieces were grown for 10 days in continuous light and transferred to a dark environment for vertical growth for two weeks (N=20) (A) and four weeks (N=55) (B). Boxes represent data points between the 25th and 75th percentile, the whiskers correspond to the minimum, and maximum measurements and lines inside the boxes represent the mean. Significance was evaluated by a 2-tailed Student’s t-test of the mutant compared to wild-type. In addition to the modest growth phenotypes, there was a clear distinction between the wild-type and ΔPpEG16 plants in the initiation of senescence. Maturation of the P. patens typically 105 proceeds by expansion, and eventually, the oldest, central gametophores senesce, becoming small and brown (Mukae et al., 2015). Scoring this trait at six weeks of growth showed that out of 36 plants, 75% and 80% of ΔPpEG16-1 and ΔPpEG16-2 plants, respectively, had a brown center versus 13% of wild-type plants (Figure 4.3). This phenotype indicates that the PpEG16 knockout plants display premature senescence of gametophytes compared to wild-type. Figure 4.3. Assessment of senescence of ΔPpEG16 and wild-type plants. (A). Representative images of senescing phenotype. Scale bars represent 2 mm. (B). Quantification of plants with browning centers after six weeks of growth on minimal media. 4.3.3 PpEG16 localizes to the cell wall at the tips of protonema Although promoter-GUS analysis has indicated the tissue-specific distribution of PpEG16 (Yokoyama et al., 2010), the subcellular localization of EG16 members was unknown. Curiously, most EG16 genes, including PpEG16, do not encode a signal peptide (Behar et al., 2018), which 106 would suggest cytosolic localization, yet the apparent substrates of PpEG16, MLG, and xyloglucan (and possibly amorphous cellulose) are cell-wall associated polysaccharides (Chapter 3; Figure B.4). Unfortunately, chloroplast autofluorescence precluded direct visualization of PpEG16-mCherry by live-imaging confocal fluorescence microscopy (Figure C.5). There was also some red autofluorescence in the gametophyte cell wall of wild-type plants (Figure C.5). To overcome this problem, immunofluorescence experiments were performed using anti-mCherry antibodies and a green (Alexa Fluor 488-tagged) secondary antibody. In fixed cells, the fluorescent label of PpEG16 was localized to the cell wall of protonema cells expressing the PpEG16-mCherry construct (Figure 4.4, Figure C.7), which was absent in the wild-type and negative controls (Figure C.6). Staining of the lateral parts of the cell walls was also observed in line PpEG16-mCherry 2-10 (Figure C.6). Faint diffuse staining of the cytosol was also seen in mCherry lines. Intracellular puncta were observed in both the transgenic and wild-type plants, suggesting non-specific fluorescence (Figure 4.4, Figure C.6). Taken together, these results suggest that PpEG16 is apoplastic, despite the absence of a canonical secretion signal peptide. 107 Figure 4.4. Confocal microscopy of PpEG16-mCherry localization in cell walls of protonema cell tips. Whole-mount immunostaining was carried out on fixed 7-day-old wild-type or PpEG16-mCherry expressing protonema using anti-mCherry and anti-xyloglucan antibodies. Wild-type cells not expressing the epitope-tagged PpEG16 did not label. Scale bars represent 50 µm. TD – transmitted light differential interference contrast images. 108 The observation that PpEG16 exhibits the highest activity on MLG (Chapter 3; Figure 3.3; Error! Reference source not found.) presents an additional conundrum, as this polysaccharide has only been observed in grasses (Poaceae), horsetails (Equisetaceae), and certain green algae (Popper et al., 2011). No antibody label was detected in the cell walls of P. patens wild-type and ΔPpEG16-1 plants using anti-MLG (BG1/BS 400‐3) antibodies (Figure 4.5). This outcome is consistent with previous analyses that indicated that MLG is absent in moss species, including P. patens (Popper and Fry, 2003, Moller et al., 2007). Figure 4.5. Confocal immunofluorescence microscopy of protonema treated with anti-xyloglucan antibodies (LM15) and anti-MLG antibodies (BG1). TBST treated tissues serve as negative controls. Scale bars represent 50 µm. TD – transmitted light differential interference contrast images. 109 In addition to MLG, PpEG16 has activity on xyloglucan (Chapter 3; Figure B.4), which has been reported in mosses (Roberts et al., 2012). Immunofluorescence of xyloglucan using the LM15 antibody, which targets XXXG-type non-fucosylated xyloglucans (Vincken et al., 1997, Marcus et al., 2008), revealed wall-specific labelling of wild-type and ΔPpEG16-1 protonema. The brightest fluorescence was observed at the cell tips and the intracellular compartments of the cell (Figure 4.4, Figure 4.5), which is similar to PpEG16 localization (Figure 4.4). There were no apparent differences in labelling between wild-type and ΔPpEG16-1 tissues (Figure 4.5). Previous studies using LM15 showed that 1- and 4-day-old P. patens protoplasts display weak staining of the cell walls (Berry et al., 2016), likely due to the difference in tissue and growth media used. Although XXGG-type xyloglucans are predominant in mosses (Schultink et al., 2014, Berry et al., 2016), XXXG-type xyloglucan has been demonstrated in P. patens tissue, particularly in ammonium tartrate-supplemented media grown protonema (Moller et al., 2007). 4.4 Discussion Although there is a great deal of knowledge on the EG16 gene distribution and frequency in plant genomes, biochemical characteristics, and structure, it is not known what the biological purpose EG16 orthologs serve. Reverse genetics experiments in Physcomitrella suggest EG16 plays a role in modulating growth and development, although the molecular mechanisms underlying this remain elusive. PpEG16 deletion plants display enhanced growth compared to wild-type plants, particularly in the earlier stages of colony growth (Figure 4.1, Figure 4.2). This result is counter-intuitive, as other GH16s, XTHs, and endo-glucanases, are implicated in the elongation of plants (Rose et al., 2002). If EG16 gene products are hydrolyzing hemicelluloses in the cell wall, then loss of these hydrolases is predicted to decrease wall loosening and lead to 110 smaller cells. Conversely, larger plants were found when XTHs or endo-1,4-β-glucanases were overexpressed (Park et al., 2004, Shani et al., 2004). However, it may be that the lack of EG16 is allowing more access to the substrate for other hydrolases to act on, or the lack of EG16 is causing the plant to compensate with higher expression of other hydrolases. In addition to the growth phenotype, ΔPpEG16 plants exhibited earlier senescence than wild-type plants (Figure 4.3). This outcome may be due to the larger size of the PpEG16 deletion plants. Localization of EG16 to the cell wall in the tips of growing protonema cells suggests that this member of the GH16 family is actively involved in modifying the cell wall. All EG16 genes discovered thus far lack a signal peptide besides several gymnosperm homologs (Behar et al., 2018), but their target substrate is present in the Golgi apparatus and the cell walls (Eklöf et al., 2013). It is possible for proteins lacking a signal peptide to be secreted (Agrawal et al., 2010, Davis et al., 2016). In fact, a recent report demonstrated that a leaderless XTH gene product from A. thaliana is exported to the cell wall by exocyst-positive organelles (De Caroli et al., 2021). P. patens transgenic plants containing a copy of PpEG16 with a C-terminal mCherry tag were used to determine subcellular localization. Live-imaging of PpEG16-mCherry tissues was not successful, as autofluorescence, possible low expression, or a combination of the two prevented the ability to discern localization (Figure C.5). However, immunolocalization showed that anti-mCherry treated protonema tissue was stained at the cell tips at the cell walls (Figure 4.4, Figure C.6). This result suggests that PpEG16 is secreted in a signal-sequence independent pathway, opening up a further study topic. Localization to the cell wall places EG16 in the same compartment as potential cell wall polysaccharide substrates. PpEG16 has hydrolytic activity on MLG, with low activity against xyloglucan and soluble cellulosic substrates (Chapter 3; Figure B.4). Although MLG labelling 111 was not found, xyloglucan immunostaining shows binding of the cell walls, appearing brighter at the tips (Figure 4.5). The co-localization of xyloglucan and PpEG16 suggests a possible interaction and could be influencing substrate availability for other hydrolases such as XTHs, which are known to be involved in cell growth (Hayashi and Kaida, 2011). Although PpEG16 activity in vitro was strongest for MLG (Chapter 3; Figure B.4), MLG staining of protonema tissues was similar to background controls (Figure 4.5). In comparison, MLG immunolabelling in maize (Zea mays) and horsetail (Equisetum arvense) is continuous throughout the cell walls (Sørensen et al., 2008, Kozlova et al., 2020). While this may be due to a low amount of MLG in the cells, or the epitopes may be masked, it is consistent with previous reports using BG1 antibodies on P. patens tissues (Moller et al., 2007). The question of why an MLGase is highly expressed and present in cell walls lacking its primary substrate is unresolved. The main activity of EG16s may be a vestigial feature, and the relatively low activity on xyloglucan and soluble cellulose components could be an important characteristic. Further, it may be that the primary substrate of PpEG16 has not yet been demonstrated. For example, a potential substrate recently described from P. patens is a linear polysaccharide of β(1,4)-linked glucosyl residues with (1,3)-linked arabinofuranosyl residues, which is similar in structure to MLG (Roberts et al., 2018). In summary, the analyses performed in this paper are the first to expand knowledge on the biological role of EG16s, demonstrating that PpEG16 is involved in the modulation of moss growth and senescence. The localization of PpEG16 to the tips of protonema cells, where it co-localizes with one of its substrates, xyloglucan, indicates that it is a cell wall protein despite its lack of canonical N-terminal signal sequence. How hydrolysis of xyloglucan in the cell walls could be linked to a constraint of growth or control of senescence remains to be determined. It will be 112 valuable to investigate whether the subcellular localization of EG16s is conserved in distantly related plant species and the unconventional secretion mechanisms involved. 113 Chapter 5: Functional analysis of Populus endo-glucanase 16 (EG16) 5.1 Introduction Plant cells are enclosed by walls made up of a complex matrix of partially crystalline cellulose, pectin, hemicellulose, structural proteins, and in cells with secondary cell walls, lignin (Schultink et al., 2014). Hemicelluloses provide extensibility by crosslinking and tethering the cellulose microfibrils (Scheller and Ulvskov, 2010, Braybrook and Jönsson, 2016). Xyloglucan is a hemicellulose ubiquitous in land plants, making up as much as 20% of the primary cell wall dry weight (Eklöf et al., 2012). Xyloglucan is prevalent in type I cell walls, the most common primary cell wall type among angiosperms (Kozlova et al., 2020). Whereas Type II cell walls describe the walls of graminaceous monocots (Poales), in which the prominent hemicelluloses are glucuronoarabinoxylans and mixed-linkage β-glucan (MLG) (Popper et al., 2011, Kozlova et al., 2020). The cell wall architecture is produced, maintained, and modified by carbohydrate-active enzymes (CAZymes), which are essential and abundant in plant genomes (Coutinho et al., 2003). CAZymes are responsible for diverse tasks, tied not only to the cell wall polysaccharide synthesis but also to development, signalling, energy, and defence (Pinard et al., 2015). A major plant CAZyme group is the xyloglucan endotransglycosylase/hydrolase (XTH) (EC 2.4.1.207 and EC 3.2.1.151), members of which modify xyloglucan and have been implicated in various processes such as cell wall loosening and strengthening, growth, and fruit ripening (Miedes and Lorences, 2009, Zhang et al., 2017). XTH gene products are glycoside hydrolase family 16 (GH16) enzymes, a group that comprises members that are active on β(1,4) or β(1,3) linkages of glucans and galactans (Michel et al., 2001, Viborg et al., 2019). Until recently, XTH gene products were the only known GH16 114 members in plants, with bacterial mixed-linkage endo-glucanases (MLGases, also known as licheninases) recognized as a sister clade according to phylogeny models (Michel et al., 2001, Eklöf and Brumer, 2010). A study showed that a subset of putative XTH sequences was missing the characteristic C-terminal extension (PFAM XET_C, PF06955), a feature of all XTH gene products (Eklöf and Brumer, 2010). Homologs of the newly discovered group, designated as endo-glucanase 16 (EG16), were found once per plant genome and were missing from many taxa, eudicots in particular (e.g., Arabidopsis spp., Solanaceae species) (Eklöf et al., 2013, Behar et al., 2018). In contrast, XTH orthologs are found in groups of 20-60 in the genomes of all land plants. Notably, a defining feature that sets EG16 gene products apart is their broad specificity toward MLG, xyloglucan, and soluble cellulosic substrates (Eklöf et al., 2013, McGregor et al., 2017) (Chapter 3). Novel phylogenetic analyses of green algae genomes show that EG16 orthologs are more commonly observed than XTH orthologs, supporting the hypothesis (Eklöf et al., 2013) that an EG16-like group is ancestral to extant EG16 and XTH genes (Jiao et al., 2020). Comprehensive genomic analyses show that the sequences of EG16 orthologs are highly conserved, with upwards of 70% sequence identity in disparate species (Behar et al., 2018) (Chapter 3; Figure 3.1). Furthermore, the biochemical characterization of EG16 gene products shows comparable activities. Specifically, the substrate specificity and limit-digest products of poplar (Populus trichocarpa), grapevine (Vitis vinifera), and spreading earthmoss (Physcomitrella patens) EG16 enzymes are consistent, despite 500 million years since the divergence of these plant groups (Eklöf et al., 2013, McGregor et al., 2017) (Chapter 3). Recent in vivo work of the P. patens EG16 homolog demonstrated that it is highly expressed in young developing tissues, that it localizes to the cell wall in protonemal tips, and that EG16-deletion plants are larger than wild-115 type plants (Chapter 4). However, it is not yet known if these features are consistent across all EG16s. In this study, we used hybrid poplar Populus alba x P. grandidentata (P39) to research EG16 expression, subcellular localization, and phenotypes of EG16-downregulated P39 plants to analyze the previously in vitro characterized PtEG16 (Eklöf et al., 2013). We show that Populus EG16 expresses in young, expanding tissues, that Populus EG16 gene products localize to the cell wall, and that EG16-downregulation in P39 hybrid poplar does not affect growth. 5.2 Materials and methods 5.2.1 Data sources PtEG16 sequence was accessed from JGI Phytozome (accession number: poptr_0002s15460) (Goodstein et al., 2012). Microarray and RNA-sequencing expression data were acquired from the eFP browser (Wilkins et al., 2009) and PopGenIE database (Sjödin et al., 2009, Sundell et al., 2015), respectively (accession number: Potri.002G153200.1). 5.2.2 Poplar growth conditions Hybrid poplar Populus alba x Populus grandidentata (P39) was grown in woody plant media (WPM) media (Lloyd and McCown, 1981) supplemented with 0.01 µM 1-naphthaleneacetic acid (NAA) and Plant Preservative Mixture (PPM; Plant Cell Technology) in Magenta vessels (GA-7; Millipore Sigma) under 16-hour light to 8-hour dark cycle with full-spectrum lighting at 24°C. Individual plants were cultured using sterile technique every six months. 5.2.3 gDNA extraction 100 mg of leaf tissue was ground with a pestle in a 1.7 mL tube and subsequently mixed with 400 µL DNA extraction buffer (200 mM Tris-HCL, 250 mM NaCl, 25 mM EDTA, 0.5% 116 (w/v) SDS). The preparation was centrifuged for 3 minutes at 20,000 x g, and the supernatant was then gently mixed with an equal volume of isopropanol and left to incubate for 10 minutes at ambient temperature. After centrifugation at 20,000 x g for 5 minutes, the supernatant was discarded, and the pellet was washed with 70% ethanol, centrifuged once more as described, and then the pellet was air-dried in a sterile environment. Finally, the pellet was resuspended in 100 µL sterile ddH2O. 5.2.4 Cloning For cloning of the promoter-GUS transgenics, the promoter, a 2000 bp sequence upstream of the translational start of PtEG16 gene and flanking sequence fragments were amplified from P. trichocarpa Nisqually-1 gDNA by PCR (Table D.1). The product was inserted into pCR-Blunt II-TOPO and transformed into E. coli using heat shock and grown on LB plates containing antibiotics for selection. The 2000 bp section directly upstream of the EG16 start codon was transferred to Gateway donor vector pDONR221-Zeo using a BP reaction, and an LR reaction was then employed to transfer the promoter into pMDC162. A plasmid preparation of the final construct was then used to transform Agrobacterium tumefaciens EHA105, as previously described (Wise et al., 2006). For the RNAi (EG16-downregulated) lines (RC8), the PtEG16 coding sequence was amplified from P. trichocarpa Nisqually-1 gDNA, inserted into pCR-Blunt II-TOPO and transformed into E. coli. A segment that does not share homology to other parts of the genome was amplified (Figure D.5) from the pCR-Blunt II-TOPO plasmid, cloned into pDONR221 using a BP reaction (Gateway Cloning, ThermoFisher Scientific), and then transformed into E. coli. An LR reaction was then used to transfer the RNA fragment into pHELLSGATE12 and transformed into E. coli. Finally, the plasmid was transformed into A. tumefaciens EHA105. 117 For the overexpression lines, the coding sequence of EG16 and flanking regions were amplified from a cDNA preparation of P. trichocarpa Nisqually-1 by PCR. This fragment was cloned into pCR-Blunt II-TOPO and transformed into E. coli. The PtEG16 coding sequence was then amplified by PCR, cloned into pDONR221-Zeo using a BP reaction, and then transformed into E. coli. The coding sequence was transferred into pMDC32 using an LR reaction. The plasmid pMDC32 containing the PtEG16 coding sequence was then transformed into A. tumefaciens GV3101. 5.2.5 Plant transformation Arabidopsis thaliana plants were transformed by the flower dip method described previously (Zhang et al., 2006). P39 poplar plants were transformed by Agrobacterium-mediated transformation on leaf discs as described (Coleman et al., 2006). 5.2.6 Genotyping gDNA was extracted from leaf tissue of each plant line and frozen in liquid nitrogen as described above. PCR was carried out using 40 ng as a template (Standard Taq; NEB) (Figure D.2). PCR products were loaded onto a 1% agarose gel supplemented with SYBR Safe DNA Gel Stain (Invitrogen) and run at 125 V for 50 minutes alongside a DNA ladder (2-Log DNA Ladder; NEB). 5.2.7 RNA extraction, cDNA preparation and RT-qPCR RNA was extracted from leaf tissue as previously described (Kolosova et al., 2004), treated with DNase (DNA-free kit; Ambion) and subsequently, cDNA was prepared using the iScript cDNA Synthesis kit following manufacturer’s instruction (Bio-Rad). RT-qPCR of the RNAi lines was completed using SsoAdvanced Universal SYBR Green (Bio-Rad). cDNA was diluted to 1:5 in the samples with primers targeting PtEG16, and 1:200 for the samples containing primers to 118 amplify housekeeping gene 18S. Samples were prepared in triplicates in a 96-well plate. A two-step protocol was used on a CFX96 Touch Real-Time PCR Detection System. 5.2.8 Tissue sectioning Fresh tissue was sandwiched between two expanded polystyrene pieces for physical support and sectioned laterally using a sliding microtome (American Optical 860 Sliding Microtome) to a thickness of 80-120 µm. 5.2.9 Promoter-GUS analysis Six GUS-lines were grown in the greenhouse for three months, after which stems, petioles, and leaves were collected from the 2nd and 8th node and then sectioned. Sections were immersed in 90% cold acetone and incubated at 4°C for 20 minutes, after which they were washed with rinse solution (50 mM sodium phosphate buffer pH 7.2, 1 mM K3Fe(CN)6, 0.5 mM K4Fe(CN)6). Staining Solution (0.1 M sodium phosphate buffer pH 7.2, 10 mM EDTA, 1 mM K3Fe(CN)6, 0.5 mM K4Fe(CN)6, 1 mM X-Gluc) was applied next, and the tubes were vacuum infiltrated three times for 1 minute each in a desiccator after which the tissue was incubated for 1 hour at 37°C to allow for the GUS enzyme to react with the substrate. For chlorophyll removal, sections were rinsed in ethanol solutions of increasing concentrations (15% to 100%), then gradually decreasing to 70% ethanol. 5.2.10 Antibodies Custom polyclonal antibodies targeting the full-length recombinant PtEG16, produced in E. coli as previously described (Eklöf et al., 2013), were generated by Cedar Lane (Burlington, Ontario, Canada). Additional anti-PtEG16 polyclonal antibodies were raised against a polypeptide sequence, VEKREGEGFPEKP, that is unique to PtEG16 and other Populus species and distinct from P. trichocarpa XTH gene products (Agrisera, Vännäs, Sweden). 119 5.2.11 Western blot PtEG16 was serially diluted (1 - 0.0001 µg) and boiled with Laemmli sample buffer (60 mM Tris-HCL, 1.5% (w/v) SDS, 8% (w/v) glycerol, 0.005% bromophenol blue, 100 mM dithiothreitol) for 15 minutes at 95°C. The samples were then loaded twice into different SDS-PAGE gels (Bio-Rad 4–20% precast protein gels) alongside a protein ladder (BLUelf prestained protein ladder) and run at 180 V for 45 minutes. One gel was stained with Coomassie dye for 20 minutes, destained overnight, and imaged (Bio-Rad ChemiDoc MP Imaging System). The other gel was transferred to a PVDF membrane (Millipore-Sigma) in a Bio-Rad Trans-Blot® SD Semi-Dry Transfer Cell in Towbin transfer buffer (25 mM Tris, 192 mM glycine, 10% (v/v) methanol) sandwiched between two thick pieces of filter paper (1703958; Bio-Rad) at 25 V, 0.35 mA, 5w for 50 minutes and then left to dry overnight. Next, the membrane was activated with 100% methanol and incubated with blocking buffer (5% (w/v) skim milk powder in Tris-buffered saline, 0.1% Tween-20 detergent (TBST)) for 1 hour at ambient temperature with gentle agitation. The membrane was then incubated with primary antibody, diluted 1:1000 in TBST, for 1 hour with continuous rocking at room temperature. The membrane was then washed with TBST for 10 minutes each time and then incubated with 1:10,000 diluted, HRP-conjugated secondary antibody (AS10 833; Agrisera) for 1 hour with continuous rocking at room temperature. Next, the membrane was washed three times for 10 minutes each. Lastly, a western substrate kit (CCH345; Bio-Helix) was used following the manufacturer’s instructions and imaged (Bio-Rad ChemiDoc MP Imaging System). 5.2.12 Immunofluorescence Tissue fixation protocol was adapted from (Zachgo et al., 2000). Stem, petiole, and leaf samples isolated from the 1st and 8th node, were sectioned as described above and incubated in 120 fresh fixative solution (50mM PIPES pH 7.5, 5 mM MgSO4, 5 mM EGTA pH 7.5, 4% (w/v) paraformaldehyde) for 1.5 hours with gentle agitation, and then washed twice in TBST buffer (10mM Tris pH 7, 0.25M NaCl, 0.1% Tween-20). Fixed samples were bleached in ethanol solutions by gradually increasing concentrations for 10 minutes each with slight agitation (25%, 50%, and 100%), and then they were once more incubated with the solutions in reducing concentrations. TBST was then used to wash the samples for 10 minutes. In 1.7 mL tubes, the samples were incubated in blocking solution (5% bovine serum albumin in TBST) for 1 hour at room temperature with gentle agitation. Subsequently, the blocking solution was discarded and incubated with primary antibody diluted in TBST (1:16 anti-PtEG16(full-length) antibodies, 1:20 anti-PtEG16(peptide) antibodies, 1:36 anti-homogalacturonan antibodies (JIM7, CarboSource, USA), 1:36 anti-xyloglucan (LM15, ab190146; Abcam), 1:36 anti-(1,3;1,4)-β-glucan (anti-MLG) (BG1/BS 400‐3, Biosupplies Australia)) for 2 hours at room temperature. The samples were then washed twice with 200 µL TBST for 5 minutes and then incubated with secondary antibody (Alexa Fluor 488 anti-rat (A-11006), anti-mouse (A-11001) or anti-rabbit (A-11008) from Invitrogen) overnight at 4°C with gentle agitation. Finally, the samples were washed twice as described above. 5.2.13 Microscopy GUS samples were mounted in 70% glycerol on microscope slides (FroggaBio, Concord, Ontario, Canada), covered with coverslips, imaged with Leica DMR microscope, and captured with a Canon EOS Rebel T5 camera using Leica 10x/Air, 40x/Air and 64x/oil lenses. Immunostained samples were mounted on microscope slides in TBST and covered with coverslips. The samples were imaged on an Olympus FV1000 Laser Scanning microscope using 10x/NA 0.4/Air, 40x/NA 0.95/Air, and 60x/NA 1.1; WD 1.5mm lenses with Alexa Fluor 488 default settings. 121 5.2.14 Growth assay Wild-type and several RNAi lines (RC8-4, RC8-6, RC8-8, RC8-10, RC8-14 and RC8-16) were grown up for five weeks in tissue culture and then potted in perennial soil (50% peat, 25% hummus, 25% crushed bark chips) in 2-gallon pots in a greenhouse. The photoperiod length was 16 hours, and the LED light output was ca. 250 µmol/m2/s, at the table surface. After 4.5 months of growth, the following parameters were measured: plant height, number of leaves, 3rd petiole and 8th petiole diameter at the base, 3rd and 8th petiole total length, 3rd-4th and 8th-9th internode stem diameter (below top petiole), and 3rd-4th and 8th-9th internode stem length. Data were first analyzed with one-way ANOVA and post hoc Student’s t-tests. Bonferroni-Holm correction was applied to minimize the problem of multiple testing. 5.2.15 Soluble sugar analysis As part of phenotyping the RNAi plants, soluble sugars were quantified in the leaves of 3-months-old RNAi and wild-type plants. To determine a difference between the higher expressing tissues of EG16 and lower expression, first to fifth leaves (designated as “young”) and eighth leaf (designated as “mature”) were collected from greenhouse-grown plants and frozen in liquid nitrogen. The analysis was carried out closely following a previously described protocol (Da Ros et al., 2020). The data were analyzed with one-way ANOVA and post hoc Student’s t-tests with Bonferroni-Holm correction. 5.2.16 MALDI-imaging Specimens (3rd – 4th internode stems) were fixed in 4% formaldehyde in PBS. Samples were vacuum dried and stored at 4°C. After fixation, the samples were embedded in 10% low melt agarose. Agarose blocks were incubated at 4°C overnight and covered with wet tissue paper to prevent dehydration during hardening. Agarose-embedded specimens were then cut in 80 µm thick 122 sections using a vibratome (Leica VT 1000S). Sections were collected and stored in 70% ethanol at room temperature. The sections were subjected to an alcohol-insoluble residue (AIR) preparation to eliminate possible protein or lipid contamination. The AIR preparation comprises three main incubation steps, 2 minutes each: 70% aqueous ethanol; 1:1 (v/v) methanol-chloroform; absolute ethanol. After each step, the eluant was removed and the sample air-dried for 1 minute. AIR sections were then stored in 70% ethanol at room temperature. Indium-tin-oxide (ITO) coated slides (Bruker, Bremen, Germany) for MALDI-Imaging were poly-L-lysinated (poly-L-lysine (PLL) solution; P8920; Sigma). Sections were placed on these PLL-ITO-coated slides. Images of sections were acquired with a bright-field microscope (Leica Application Suite, Leica camera DFC 420C). The images of sections were pasted together and scaled using ImageJ software (Preibisch et al., 2009, Schindelin et al., 2012). The enzyme solutions were sprayed with an HTX TM-sprayer (Bruker Daltonic) at 30°C on sections placed on the PLL-ITO-coated slides. Xyloglucan was enzymatically digested with 50 mu/µl endo-β-1,4-glucanase (XEG) (Pauly et al., 1999) in 50 mM ammonium formate buffer, pH 4.5. MLG was treated with 50 mu/µl of MLGase (Megazyme, resuspended in 50 mM ammonium acetate, pH 6.5). Both enzyme solutions were sprayed at a flow rate of 7.5 µl/min and 660 mm/min. As a positive control to verify enzyme digestion, 1 µl of A. thaliana AIR (10 mg/mL) and 1 µl of barley AIR (10 mg/mL) were spotted on the slides. After spraying the enzyme, the slide was incubated for 20 minutes at 37°C and 1 hour at 50°C. The matrix chemical used for MALDI-TOF analysis is 100 mg/mL of 2,5-dihydroxybenzoic acid (DHB) dissolved in a mixture of 1:1 H2O:Acetonitrile (Carl-Roth) containing 2% of N,N-dimethylaniline (DMA, Fisher Scientific) (Snovida et al., 2008) and 15 mM 123 NaCl. The matrix was sprayed on the (digested) samples using the HTX TM-sprayer at 25°C, at a flow rate of 0.1 mL/min. As controls, to assess technical aspects of this method, 1 µl β-1-3-cellobiose (10 mg/mL) solution was spotted onto the MLG digested sample on the slide or 1 µl of a mixture of tamarind xyloglucan oligos (Megazyme) on the xyloglucan digested samples. Both solutions were spotted before matrix deposition and vacuum dried. The analysis was performed with a Rapiflex Tissuetyper MALDI-TOF (Bruker, Bremen, Germany) controlled with FlexControl 4.0 (Bruker, Bremen, Germany) in positive reflector mode collecting 2000 hits per laser shot. The laser settings were adjusted to 20 µm laser spots and 50 µm laser diameter. The mass/charge range for xyloglucan digestion was 600-3200 m/z, for MLG digestion 400-3200 m/z. An optical image of the ITO slide was obtained using the FlexImaging 5.0 (Bruker, Bremen, Germany) software for selected areas. The raster width for obtaining the images was set to 25x25 µm. The images were processed using FlexImaging 5.0. The resulting spectra were normalized according to the root mean square (RMS) of the data points (Deininger et al., 2011). 5.3 Results 5.3.1 PtEG16 expression is highest in young and expanding tissues The eFP browser and PopGenIE databases were queried to determine the expression of EG16 in various P. trichocarpa tissues (Figure D.1). Expression was highest in male (584.36 ± 108.95 RMA) and female catkins (517.4 ± 123.61 RMA) (Figure D.1A). Young leaves and roots follow with two-fold lower expression, and lastly, seedlings and mature leaves expressing the lowest levels, ca. four-fold and 10-fold lower than that of young leaves, respectively. Consistent with the microarray data, RNA-sequencing data shows that the highest expression is in expanding 124 catkins and field-grown expanding leaf buds and the lowest expression in mature leaves (Figure D.1B). To discern expression differences between various tissue types and cells, hybrid poplar P39 was transformed with a PtEG16 promoter::β-glucuronidase (pPtEG16::GUS) construct (Figure D.2A). Informed by the expression data from microarray and RNA-sequencing studies, a promoter-GUS analysis was carried out with contrasting young and mature tissue (Figure 5.1). GUS staining of young and mature leaves, petioles, and stems showed that expression is strong in the epidermis, cortical parenchymal cells, and in the pith of young (2nd node) petioles and stems, and is almost non-existent in mature tissues (8th node) (Figure 5.1). Young leaves showed higher expression in the apical side of the blade, in the epidermis and cortex, and lower expression in the epidermis of the leaf vein. No staining was observed in the cortex, pith, and vascular tissues of the vein. GUS-stained young stems and petioles presented a pattern in which staining was pronounced in cells surrounding sclerenchyma cells in the cortex (Figure 5.1). 125 Figure 5.1. Promoter-β-glucuronidase assays of transverse sections of P39 poplar lines expressing pPtEG16::GUS. Scale bars represent 0.1 mm. E – epidermis; Co – cortex; C – cortical collenchyma cells; P – cortical parenchyma cells; S – sclerenchyma; Ph – phloem; V – vascular cambium; X – xylem; Pi – pith. 126 5.3.2 Populus EG16 immunolocalization Subcellular localization is imperative to gain further insight into the biological function of the EG16 gene product. As EG16s do not contain signal peptides as part of their sequence, cytosolic localization would be predicted. However, that would not be consistent with the localization of the substrates of EG16 enzymes, MLG, xyloglucan, and cellulosic substrates, which are present in the Golgi bodies and cell walls (Eklöf et al., 2013). For localization, antibodies were custom-made for the full-length PtEG16 protein. Immunolocalization experiments using the anti-PtEG16(full-length) antibody showed binding of the cell walls of all tissues used (Figure D.4). However, the binding to tomato (Solanum lycopersicum) cell walls suggests non-specific binding, as the plant does not have an EG16 homolog (Behar et al., 2018). Moreover, the strong fluorescent signal from mature petioles contradicts EG16 expression analyses that showed low expression in older tissues (Figure 5.1). To avoid non-specific binding, peptide-specific antibodies were custom-made to target a PtEG16 peptide that is not present in PtXTH orthologs or other proteins in the P. trichocarpa proteome (Figure D.3). The binding of the antibodies was demonstrated with recombinant PtEG16 produced in E. coli (Figure D.5). Immunofluorescence experiments using the peptide-specific antibody on wild-type and EG16-downregulated petiole cross-sections showed a stronger signal in young, wild-type tissues (Figure 5.2). The binding observed in young petiole sections corresponds to the cell walls of the cortex, with some binding to the cell walls of vascular tissue. In mature wild-type tissues, the signal appeared as large intracellular puncta, while in the EG16-downregulated tissues, faint binding to vascular cell walls was observed, although this may be due to autofluorescence (Figure 5.2). Contrary to expectations from promoter-GUS analysis, a signal also appears in the cortical collenchyma cells of young petioles where EG16 expression is low 127 (Figure 5.2). Generally, these observations indicate specific binding, as strong binding corresponds to tissues where EG16 is expressed and weak or no binding in downregulated and mature tissues. 5.3.3 Populus EG16 downregulation does not affect plant growth PtEG16 expression was downregulated in P39 hybrid poplar by RNA interference (RNAi). Of the 20 lines selected, 15 contained the construct after PCR verification (Figure D.2B). The RT-qPCR analysis showed that expression was downregulated by 60-90% in all lines except for lines 11 and 13 (Figure D.6). The lines RC8-4, RC8-6, RC8-8, RC8-10, RC8-14 and RC8-16 were subsequently selected and planted in the greenhouse for 4.5 months. Morphological differences were not observed between the wild-type and RNAi plants when inspecting numerous parameters over the growth period. Specifically, plant height, number of leaves, young and mature petiole diameter and length, and young and mature internode stem diameter and length were measured (Figure 5.3). Line RC8-4 was the exception of the six lines, where the plant had significantly larger young and mature petiole diameter measuring 119% and 121% compared to wild-type, respectively, and longer mature petioles, measuring 122% longer than wild-type (Figure 5.3). As only one line of the six presented with growth phenotypes, it appears that EG16-downregulation does not affect plant growth in the conditions and tissues tested. In addition to the downregulation of PtEG16, constructs were made to overexpress PtEG16 under a 35S constitutive promoter in Arabidopsis thaliana. However, two repetitions of the transformation procedure did not produce viable transformants (data not shown). 128 Figure 5.2. Confocal microscopy of immunolabelled P39 poplar (A) and EG16-downregulated (RC8-6) (B) sections. Sectioned petiole tissue from greenhouse-grown plants. Young – 2nd node petiole, mature – 8th node. Scale bars represent 100 µm. E – epidermis; C – cortical collenchyma cells; P – cortical parenchyma cells; S – sclerenchyma; Ph – phloem; X – xylem. 129 130 Figure 5.3. Phenotyping of EG16-downregulated lines. Growth parameters of each line (N = 8) were measured after 4.5 months of growth. Boxplots represent measurements from 25th to 75th percentile, and whiskers denote the minimum and maximum measurements. 5.3.4 Analysis of PtEG16 substrates in P39 tissues To determine if the downregulation of PtEG16 affects its substrates, MLG presence and xyloglucan localization in situ were analyzed. As previously described, MLG is the prominent substrate of EG16 gene products (McGregor et al., 2017) (Chapter 3) and PtEG16 in particular (Eklöf et al., 2013). However, MLG has not been identified in poplar tissues thus far (Popper et al., 2011). Two approached were taken to determine whether MLG may be present in EG16-expressing tissues, immunostaining and MALDI-imaging. For immunostaining, anti-MLG antibodies were used to treat young and mature petioles of wild-type and EG16-downregulated plants. Staining appeared as dim puncta covering various areas of the tissue (Figure D.7A). In contrast, immunostaining of plants that have MLG, display continuous fluorescence in the cell walls (Kozlova et al., 2020). Therefore, the observed pattern is likely unspecific binding. In addition, MALDI-imaging was used as an alternative method to determine the presence of MLG ions in stems of wild-type and EG16-downregulated plants. The undigested control and the MLGase-digested samples did not display MLG-related ions, namely cellotriose (GGG, m/z 527) or cellotetraose (GGGG, m/z 689), in both wild-type and RNAi lines (Figure D.7B). Hence, no MLG was detected using two methods under the specified conditions. Xyloglucan is prominent in poplar species as in other eudicots (Pauly et al., 2013) and is a secondary substrate of PtEG16 (Eklöf et al., 2013). To determine whether there may be differences in xyloglucan oligosaccharide distribution between wild-type and EG16-downregulated plants, tissues were treated with anti-xyloglucan antibodies to determine possible differences in 131 localization and cell shape in wild-type and EG16-downregulated petioles. All tissues tested displayed smooth lines surrounding every cell without meaningful differences between genotypes (Figure 5.4A-B). In addition, MALDI-imaging was done using a xyloglucan endo-β-1,4-glucanase (XEG). The epidermis, phloem, and collenchyma tissues of young stems were probed, and m/z ratios of xyloglucan degradation ions were inspected. The expected eudicot xyloglucan oligosaccharides were observed in all samples, and their distribution did not differ between the plant lines and the tissues tested (Figure D.8). Imaging the entire stem sections showed that the pith lacked xyloglucan ions in all samples, likely due to the presence of large cells at low density in this region (Figure 5.4C-D). Additionally, imaging shows that the four xyloglucan ions visualized are present in similar proportions to each other in all tested samples. Xyloglucan was observed in all tissue regions, concordant with immunostaining experiments (cf. Figure 5.4A-B and Figure 5.4C-D). To further probe differences between the wild-type and RNAi lines, the soluble sugars galactose, sucrose, myo-inositol, fructose, raffinose and glucose were quantified in young and mature leaf tissue collected from 3-month-old greenhouse plants. Generally, soluble sugar quantities did not consistently differ from wild-type (Figure D.9). Lines RC8-8 and RC8-16 tissues had significantly lower glucose than the wild-type, with 0.34 ± 0.04 µg·mg-1 (p-value = 1·10-4) and 0.57 ± 0.01 µg·mg-1 (p-value = 4·10-6), respectively in young tissues and 0.25 ± 0.02 µg·mg-1 (p-value = 3·10-4) and 0.52 ± 0.01 µg·mg-1 (p-value = 1·10-4), respectively, in mature tissues. However, these disparities were not consistent across all the downregulated lines (Figure D.9). 132 Figure 5.4. Analysis of PtEG16 substrate xyloglucan in P39 tissues. Confocal microscopy of immunolabelled P39 poplar (A) and EG16-downregulated P39 (B) using anti-xyloglucan antibodies (LM15). Scale bars represent 100 µm. E – epidermis; C – cortical collenchyma cells; P – cortical parenchyma cells; S – sclerenchyma; Ph – phloem; X – xylem. MALDI-imaging of xyloglucan ions in the tissues of P39 poplar wild-type (C) and in two EG16-downregulated lines (D). Sections of 3rd-4th internode stems were treated with a xyloglucanase and inserted into a MALDI- TOF mass spectrometer. Images of a selected ion were assembled from the mass spectrum obtained from each 5 µm spot. 133 5.4 Discussion A 2013 study describing the biochemical and phylogenetic characterization of PtEG16 was the first to distinguish EG16 gene products as a distinct clade of plant GH16 enzymes vis-à-vis the well-known XTH gene products (Eklöf et al., 2013). Since then, an extensive bioinformatic analysis of EG16 orthologs was performed to discern the EG16 gene evolutionary origin, distribution, and frequency in the plant kingdom (Behar et al., 2018). Additionally, two EG16 orthologs from V. vinifera and P. patens were characterized biochemically, including determining enzyme structure by X-ray crystallography (McGregor et al., 2017) (Chapter 3). In P. patens, the expression, localization, and growth phenotype of EG16-deletion plants has been demonstrated (Chapter 4). This chapter demonstrates that, as with the conservation of PtEG16 and PpEG16 protein sequences and enzymatic activity, PtEG16 and PpEG16 gene expression patterns and localization are analogous, suggesting a similar biological function. Promoter-GUS assays in P39 hybrid poplar show that EG16 expression is high in young tissues, specifically in cortical parenchyma cells, with little expression apparent in cortical collenchyma cells and vascular bundles (Figure 5.1). Therefore, similar to P. patens EG16 and other EG16 orthologs, PtEG16 expression is associated with young and rapidly growing tissues and cell types (Chapter 4). The contrast in expression between the two cortical cell types is striking. Collenchyma cells have thick primary cell walls made up of large amounts of cellulose and small amounts of hemicellulose and pectin and are present in fast-growing tissue, serving as mechanical support (Crang et al., 2018). In contrast, cortical parenchyma cells are metabolically active, photosynthesizing and secreting cells (Crang et al., 2018). It may be that the reason for the difference in expression is that EG16 is active in cells that are growing, dividing, and also have higher amounts of xyloglucan. The association of EG16 expression with young tissues in 134 bryophyte, gymnosperm, and eudicot species suggests a conserved role in tissue development (Behar et al., 2018). As EG16 genes do not encode signal peptides, it is essential to determine localization to understand fully EG16 gene product function in vivo. Observations from immunolocalization experiments showed that EG16 localizes to cell walls (Figure 5.2). This result is consistent with previous studies of EG16 in P. patens protonema tissues, which showed localization to the cell wall at the cell tips (Chapter 4; Figure 4.4). Though not well studied, there are secretion mechanisms that are independent of classical endoplasmic reticulum and Golgi apparatus secretion in eukaryotic cells, such as by plasma membrane blebbing or exosomes (Agrawal et al., 2010). Some plant enzymes have also been shown to be secreted unconventionally (Davis et al., 2016, De Caroli et al., 2021). Localization was observed in cells where the gene is predicted to be lowly expressed, as indicated by promoter-GUS analysis. PtEG16 promoter activity was low in cortical collenchyma cells (Figure 5.1). Nevertheless, localization using peptide-specific antibodies showed binding to both cortical collenchyma and parenchyma cell walls in young wild-type tissue (Figure 5.2). There are many indicators that the binding is specific, mainly that there is less binding observed in mature and EG16-downregulated tissues, where EG16 expression is minimal (Figure 5.1, Figure D.1). The highly sensitive immunofluorescence assay may sufficiently detect the protein produced by the low expression levels. It is also possible that the use of the 2 kb promoter in the promoter-GUS analysis is missing cis-regulatory elements required for wild-type expression. Numerous measurements showed no differences in growth between wild-type P39 hybrid poplar and the EG16-downregulated lines. Growth parameters (Figure 5.3), soluble sugars (Figure D.9), xyloglucan localization (Figure 5.4A-B), and xyloglucan oligosaccharide 135 distribution (Figure 5.4C-D, Figure D.8) did not reveal differences between wild-type plants and those low in EG16 expression. It may be that the limited amount of EG16 gene product present is adequate for performing its biological role. Alternatively, the loss may be compensated by unknown mechanisms, or EG16 may play an insignificant role in poplar development. Similarly, a knockout of a xyloglucan endohydrolase in A. thaliana did not affect plant growth and development (Kaewthai et al., 2013). In comparison, deletion of EG16 in the bryophyte P. patens produced larger plant colonies (Chapter 4). As EG16 homologs are more common in non-angiosperm plant genomes (Behar et al., 2018), they may have a more significant biological function in those species than in eudicots. In addition, the inability to produce overexpression lines in A. thaliana suggests that overexpression of EG16 may be lethal. Interestingly, the effect of EG16 downregulation in Populus may be most pronounced in catkins, the tissue that has the highest EG16 expression (Figure D.1). However, the extended time (years) required for Populus to achieve flowering maturity makes testing this hypothesis practically challenging. EG16 protein sequences are highly conserved across the plant kingdom (Behar et al., 2018), and this conservation extends to enzymatic activity, substrate specificity (Eklöf et al., 2013, McGregor et al., 2017) (Chapter 3), as well as gene expression patterns (Chapter 4; Figure C.1). The research presented here showcases the first angiosperm EG16 ortholog to be studied in vivo. We demonstrate that signal-sequence-independent localization to the cell wall is conserved, as was shown for P. patens EG16. In addition, downregulation of EG16 is not lethal to plants and was shown to either not influence the growth or yield minor growth phenotypes (Figure 5.3, Chapter 4; Figure 4.1, Figure 4.2). Overall, it remains a question why EG16 is conserved in vastly distant plant groups. Probing the precise secretion mechanism, binding to substrates in vivo, and 136 attenuated overexpression phenotypes (e.g., by using CesA6 primary cell wall promoter) may further elucidate the biological function of these enzymes. 137 Chapter 6: General conclusions and future directions 6.1 General conclusions In chapter 2, a census was conducted to elucidate the evolutionary origin, distribution, and frequency of EG16 genes in plants from different clades among the plant lineage and marine eukaryotes. The study uncovered a large number of previously unknown GH16 genes, and revealed an EG16 group distinct from the classical EG16 clade. EG16-2, distinguished by protein sequence motifs and a long N-terminus, were common in charophyte green algae and mosses, but were not observed in tracheophytes besides several leptosporangiate ferns. Classical EG16 orthologs were found mainly in non-eudicot plant clades such as bryophytes, ferns, and graminaceous monocots, but were generally absent from asterids and rosids. EG16 genes were found in a species sister to Zygnematophyceae charophyte green algae genome, K. nitens, without XTH genes suggesting that the hypothesis that an EG16-like gene is a common ancestor to current-day plant GH16 orthologs is plausible (Eklöf et al., 2013, Jiao et al., 2020). Phylogenetic analyses showed that EG16 protein sequences are incredibly conserved, and their evolutionary relationships parallel plant evolution. This observation suggests an important biological role, as selective pressures throughout land plant speciation and evolutionary development did not select for significant modifications in the sequences of EG16 orthologs. The low representation of EG16 gene-containing species in eudicot clades indicates that EG16 function(s) may be redundant or unessential in eudicot physiology. Work in chapter 3 involved the in vitro characterization of the exemplary representative EG16 ortholog from the bryophyte clade to extend the structure-function analysis of EG16 members beyond angiosperm plants. PpEG16 activity was shown to be similar to that of previously characterized EG16 gene products. PpEG16 exhibited an identical broad substrate specificity, with 138 the highest activity on MLG of the substrates tested, and with secondary activity on soluble cellulosic substrates and xyloglucan. The cleavage pattern of xyloglucan was consistent, producing comparable limit digest products with hydrolysis after the unsubstituted glucose. However, the activity was relatively low towards xyloglucan in comparison to angiosperm EG16s. The digest products from MLG hydrolysis indicated that the PpEG16 enzyme prominently targets the β(1,4) of MLG as observed for PtEG16 and VvEG16, but also acts on β(1,3) linkage to a higher degree than the other EG16 gene products. Biomechanical experiments showed that P. patens, V. vinifera, and P. trichocarpa EG16 enzymes, when applied to chia and cucumber hypocotyls, generally cause abrupt breakage of the tissue as opposed to rapid extension with acid growth or slow extension with bacterial enzyme treatment. PpEG16 did not affect cucumber hypocotyls, which could be due to the relatively lower activity of the enzyme on xyloglucan compared to VvEG16 and PtEG16. Overall, these observations indicate that EG16 enzymes are functionally conserved and can affect the cell wall structure. The research in chapters 4 and 5 sought to investigate the biological function of EG16 in two distantly related model organisms. As an extension of sequence and activity conservation, gene expression was also comparable. Moreover, in the respective tissues of P. patens and Populus, EG16 promoter expression was more pronounced in young, developing tissues. In addition, EG16 gene product subcellular localization was found to be apoplastic in both model organisms. Consistent with the conclusions from the preceding two chapters, these data suggest a conserved biological function across different plant clades. The effects of downregulation of EG16 produced different outcomes in the two model plants. EG16-downregulated Populus tissues were not phenotypically different from wild-type in their growth, cell morphology, xyloglucan deposition, and soluble sugar content. In contrast, 139 PpEG16-deletion moss plants grew larger and underwent senescence earlier than wild-type plants. This discrepancy may be attributed to the difference between downregulation and deletion, where the low EG16 expression in Populus may be sufficient for normal plant growth. It is also possible that EG16 function is redundant in angiosperm or woody plants, as suggested from the EG16 gene census of chapter 2. In addition, the larger ΔPpEG16 moss plants could have resulted from indirect higher expression of other cell wall enzymes, or EG16 gene products may have an inhibitory role in developing tissues. Although the biological function of EG16 is not entirely resolved, the data above elucidated several elements about the nature of this group of enzymes. First, EG16 genes most likely preceded XTH genes. Second, EG16 gene products from diverse species have a broad specificity towards cell wall polysaccharides, express in young tissues, localize to the cell wall, and are able to degrade plant tissues. Third, EG16 gene products may have a more substantial role in bryophyte and possibly other non-angiosperm cell walls than in those of eudicots. 6.2 Future directions As demonstrated, the EG16 clade has been characterized considerably. However, there are still many unanswered questions regarding the biology of EG16 members. EG16 biological function appears to be non-essential and possibly redundant as EG16 downregulation has minimal effects on plant physiology. However, these details contrast with the high conservation of EG16 protein sequence, enzyme function, and gene expression pattern across distant plant clades. Investigation of homologous and heterologous overexpression phenotypes may add to the understanding of the EG16 mechanism in vivo. To discern whether ΔPpEG16 plants are larger due to the lack of EG16 activity or other causes such as increased expression of genes of other cell wall modifying enzymes, RNA-seq should be performed. 140 It is unclear why the MLGase activity of EG16 gene products would be conserved in plants lacking the target polysaccharide. However, it may be that MLG is incorrectly identified as the primary substrate, and that it serves as a proxy for cellulose in the in vitro assays. If this were true, the biological targets of EG16 could instead be cellulose components. Product analysis of EG16 digests of cell wall tissue with HPAEC-PAD and MALDI-TOF, could reveal the preferred natural substrates. In addition, it is uncertain whether EG16 gene products function differently in MLG-containing cell walls of plants, such as graminaceous monocots. In vitro enzymatic characterization, protein localization, and phenotypic analysis of EG16-upregulated and downregulated transgenic plants using monocot model plants, would elucidate if they are similar to moss and eudicot EG16s. It is not presently known which secretion mechanism is employed for EG16 export to the apoplast. Since there are several pathways for the nonclassical secretion of proteins in eukaryotes (Ding et al., 2014, De Caroli et al., 2021), associated markers for each can be used to demonstrate the mechanism utilized in EG16 export. Inhibitory treatments of classical secretion pathways (e.g., brefeldin A) could be used to investigate if they affect EG16 localization, to eliminate the possibility of ER-Golgi secretion. In addition, microscopy of plant tissues co-expressing EG16 with organelle markers that label autophagosomes and exocysts, such as ATG8 and Exo70E2, respectively, could indicate if those elements are involved in EG16 targeting. The newly classified EG16-2 enzymes are not presently characterized. However, as several dozen EG16-2 genes were identified in chapter 2, the sequences could be utilized in structure-function analyses to determine whether their three-dimensional structure, substrate specificity, and other aspects are similar to canonical EG16 enzymes. 141 While the cell wall serves an essential role in plant biology, many of its inner workings remain elusive. As society greatly depends on plant growth for oxygen, nutrition, and shelter, we should seek to understand the underlying factors involved in such processes. Going forward, further investigation of the EG16 export mechanism could contribute to the understanding of the poorly studied unconventional secretion pathways. In addition, characterization of the EG16-2 enzymes could reveal unknown details about green algae cell wall biology, a currently growing field. The goals of this thesis were to characterize the newly identified EG16 family. Before the project, the understanding of EG16s depended on limited bioinformatics and in vitro biochemical characterization of two angiosperm homologs (Eklöf et al., 2013, McGregor et al., 2017). The research presented here revealed hundreds of EG16 homologs that were shown to be incredibly conserved across the plant lineages. 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Expression of PpEG16 in Physcomitrella patens tissues from the eFP browser. 162 Appendix B Supporting information for Chapter 3: Conservation of endo-glucanase 16 (EG16) activity across highly divergent plant lineages Table B.1. Primer sequences used for ligation-independent cloning into pMCSG53 Primer sequence (5->3) Variant TACTTCCAATCCAATGCCATGAATGGAGGGGATTGTGGCGGC PpEG16.1 TTATCCACTTCCAATGTTACTGATCCGGTAAATGAGGGAC PpEG16.1 TACTTCCAATCCAATGCCATGCAATGCATGGCTACTCCTAAG PpEG16.9 TTATCCACTTCCAATGTTACTGGTCGGGCAAATGGGGAAC PpEG16.9 TACTTCCAATCCAATGCCATGACGTCAAAGTGGACGATGGAT PpEG16.10 TTATCCACTTCCAATGTTATTGATCAGGAAGATGGGGAAC PpEG16.10 Table B.2. Primer sequences used for site-directed mutagenesis Primer sequence (5’->3’) Variant AAGGCGATAAGTTTCAAGATGCAATCGACTTTGAATTTCTGGG PpEG16.9 E93A, inactive mutant ATTGCGATCTCACGAGCGGCCGCGGAATGATCAAGCACCTC PpEG16.9 K34A/D35A, stability mutant GAGGTGCTTGATCATTCCGCGGCCGCTCGTGAGATCGCAAT PpEG16.9 K34A/D35A, stability mutant ATTGGCTTCATGGGGTATGCAGCAGCTGGTGCGCGTTCGTGCTTGCGAACCTG PpEG16.9 K159A/N161A/D162A, stability mutant CAGGTTCGCAAGCACGAACGCGCACCAGCTGCTGCATACCCCATGAAGCCAAT PpEG16.9 K159A/N161A/D162A, stability mutant CATGGGGTATGCATCATTTGGTGCGGCTGCGTGCTTGCGAACCTGTACCTG PpEG16.9 E157A/R158A/K159A, stability mutant CAGGTACAGGTTCGCAAGCACGCAGCCGCACCAAATGATGCATACCCCATG PpEG16.9 E157A/R158A/K159A, stability mutant TCGCAAGCACGAACGCCCAAATGATGCATACC PpEG16.9 K159 GGTATGCATCATTTGGGCGTTCGTGCTTGCGA PpEG16.9 K159, stability mutant GTATGCATCATTTGGTTTTTCGTGCTTGCGAACCTG PpEG16.9 R158, stability mutant CAGGTTCGCAAGCACGAAAAACCAAATGATGCATAC PpEG16.9 R158, stability mutant 163 Figure B.1. PpEG16 (Pp3c21_9420V3) mRNA transcripts aligned to chromosome 2. mRNA sequences acquired from JGI Phytozome (https://phytozome.jgi.doe.gov). Grey indicates untranslated regions, while teal indicates coding regions. Figure B.2. PpEG16.9 homology model. Overall model structure of PpEG16.9 in green superposed with the crystal structure of VvEG16 in grey (PDB ID: 5DZG). Active site cleft residues and catalytic residues are shown in sticks. 164 Figure B.3. Recombinant production of PpEG16 protein variants. (A). Coomassie Brilliant Blue-stained SDS-PAGE gel. (B). Intact mass spectrum of recombinant PpEG16.9. The expected mass of the full-length construct, including the 6xHis tag and TEV cleavage site, following cleavage of the N-terminal methionine, is 26373.4 Da (observed 26391.9 Da). 165 Figure B.4. Substrate screening of PpEG16 variants. The activity of PpEG16.1, PpEG16.9, and PpEG16.10 were each evaluated with 1.13 mg·mL-1 substrate and the release of reducing ends was quantified by a copper-bicinchoninic acid end-point assay. Each measurement represents an average of three replicates. Error bars reflect the standard deviation. 166 Figure B.5. Temperature (A) and pH (B) profiles of PpEG16.9 using 1 mg·mL-1 MLG. Each data point represents an average of three replicates. 167 Figure B.6. PpEG16.9 stability at 4°C using different enzyme dilution buffers. These solutions were diluted 1:10 into the assay solution, resulting in a concentration of additive (EDTA or NaCl) of 0.1 mM in the assay. 168 Figure B.7. PpEG16.9 limit digest products. HPAEC-PAD chromatograms showing the products of PpEG16.9 treatment of mixed-linkage β-glucan (A) and xyloglucan (B) after incubation for three days at optimal conditions. 169 Figure B.8. Activity of EG16 orthologs on AZCL-polysaccharides. (A). AZCL-xyloglucan incubated with 50 µg EG16 proteins at pH 5.5 and pH 7.5. (B). AZCL-HEC incubated with 50 µg EG16 proteins at pH 5.5 and pH 7.5. Error bars indicate the standard errors (n = 3). Significance was assessed using one-way ANOVA with post hoc Tukey HSD (p < 0.05). 170 Figure B.9. Matching enzyme activities of E-XEGP and VvEG16 using AZCL-conjugated HEC and xyloglucan. (A). Activity of 1 μg/mL of E-XEGP is matched by 72.5 μg/mL VvEG16 as assayed by hydrolysis of AZCL-xyloglucan. Error bars indicate the standard errors (n = 3). (B). Activity of 1 μg/mL of E-XEGP is matched by 15 μg/mL VvEG16 as assayed by hydrolysis of AZCL-HEC. 171 Figure B.10. Stress-strain analysis for EG16 orthologs normalized to xyloglucanase activity of VvEG16. Activities were normalized based on data shown in Figures S8 and S9. 3 mg of PpEG16, 297 µg PtEG16 and 2.76 µg E-XEGP were used. Error bars indicate the standard errors (n = 10). The asterisk indicates significance, as assessed by Student’s t-test (two-way, p < 0.05). Figure B.11. Effect of EDTA and DTT on the stress-strain analysis of cucumber hypocotyl walls with PpEG16. 20mM sodium acetate buffer (pH 5.5) was supplemented with 0.1 mM EDTA and 0.25 mM DTT containing 2 mg/mL of PpEG16. Error bars indicate the standard errors (n = 12). No significant difference was detected by Student’s t-test (two-way, p < 0.05). 172 Appendix C Supporting information for Chapter 4: Physcomitrella patens endo-glucanase 16 is involved in the cell wall development of young tissue Table C.1. Primers used in this study Sequence (5'-3') Purpose GTCTCTCCAGATTATTTGCCCC Amplification of 5' flanking region for producing knockout lines GTTTTACGTCTTCAGTCGCC Amplification of 5' flanking region for producing knockout lines GTGACAGAATCACGAGACATCC Amplification of 3' flanking region for producing knockout lines AGCCACAACCCACAAATCC Amplification of 3' flanking region for producing knockout lines TCACCGCGGCGAAGCTGCAAAGTGACAGAATC Amplification of 3' flanking region for producing PpEG16-mCherry lines GGATTATAATCTAGAAGCCACAACCCACAAATCCAAAC Amplification of 3' flanking region for producing PpEG16-mCherry lines GGGGACAAGTTTGTACAAAAAAGCAGGCTATGTCTCTCCAGATTATTTGCC 5'UTR of EG16 genomic sequence in frame with mCherry gene GGGGACCACTTTGTACAAGAAAGCTGGGTCCTGATCAGGGAGATGAGGA ATTB2 with EG16 sequence that will be in frame with mCherry gene TATGCGGCCGCGTCTCTCCAGATTATTTGCCCC Genotyping of EG16 deletion lines ATAGGTACCAGCCACAACCCACAAATCC Genotyping of EG16 deletion lines CACCATCGTGGAACAGTACG Genotyping primer for PpEG16-mCherry moss lines TGTGGAATTGTGAGCGGATA Genotyping primer for PpEG16-mCherry moss lines 173 Figure C.1. EG16 expression in Physcomitrella patens tissues. (A). Microarray analysis was acquired from the eFP browser (Winter et al., 2007). (B). Precise assay values. 174 Figure C.2. Genotyping of Physcomitrella patens transgenic lines. 1% agarose gels supplemented with 0.002% RedSafe of genotyping of ΔPpEG16 lines (A) and PpEG16-mCherry lines (B). (A). Primers anneal to flanking regions of the PpEG16 gene. Amplicons from the wild-type genome were expected to be 2160 bp and to include the PpEG16 gene, while amplicons from the deletion lines were expected to be 2792 bp and include the antibiotic selection cassette. (B). Bands represent the amplicons of part of the mCherry sequence of the insertion, which is expected to be 400 bp. 175 Figure C.3. Representative growth progress of wild-type and ΔPpEG16 moss lines. 176 Figure C.4. Replicate growth assays of colonies of wild-type (wt) and two lines of PpEG16-deletion mutants. 54 colonies (N = 18 per line) were measured at each time point in three independent assays. P-values, as calculated by a 2-tailed Student’s t-test, are displayed if below 5%. 177 Figure C.5. Confocal live-imaging microscopy of PpEG16-mCherry expressing line and wild-type. Excitation using 559 nm laser and emission filter of 590-640 nm was used. Scale bars represent 100 µm. TD – transmitted light differential interference contrast images. 178 Figure C.6. Whole-mount immunostaining confocal microscopy of 7-day old wild-type and PpEG16-mCherry tissues. (A). Positive and negative controls of confocal immunofluorescence microscopy of wild-type protonema. (B). Experimental images of tissue treated with anti-mCherry antibodies. Arrowheads indicated stained cell wall tips and lateral cell walls. Scale bars represent 50 µm. TD – transmitted light differential interference contrast images. 179 Figure C.7. Calcofluor-white staining of wild-type P. patens protonema tissue. Scale bars represent 100 µm. TD – transmitted light differential interference contrast images. 180 Appendix D Supporting information for Chapter 5: Functional analysis of Populus endo-glucanase 16 (EG16) Table D.1. Primers used in this study Sequence (5'-3') Purpose CAAAATGTCAAACAAAACCTACC Forward primer to amplify PtEG16 from the 5' UTR GTGTTGATTGGTCGTATTGTG Reverse primer to amplify PtEG16 from the 3' UTR GGGGACAAGTTTGTACAAAAAAGCAGGCTATGGCGGATCCAGCAATCC Forward primer to amplify PtEG16 RNAi fragment 1 with attB1 site GGGGACCACTTTGTACAAGAAAGCTGGGTTCAAGAATCACAAGAACACTCC Reverse primer to amplify PtEG16 CDS with attB2 site GGGGACCACTTTGTACAAGAAAGCTGGGTGGGATTTGTCACCTTCAA Reverse primer to amplify PtEG16 (0-256 bp CDS) with attB2 GGGGACAAGTTTGTACAAAAAAGCAGGCTCAAGACCATCGTACAAAC Forward primer to amplify PtEG16 (290-end bp CDS) RNAi fragment 2 with attB1 CAAAGCAAGGTCGAGGAGG Forward primer to clone PtEG16 promoter region from near Potri.002G153100s first exon CGTATGTGAGAGTGATGGAATTGG Reverse primer to clone out PtEG16 promoter region from within PtEG16 GGGGACCACTTTGTACAAGAAAGCTGGGTTTCTGTTTTCTTGGAAATAAATATAC Reverse primer to amplify PtEG16 promoter from topo vector with attB sites GGGGACAAGTTTGTACAAAAAAGCAGGCTGAAAGTTTTTTTTTTGAAGAAAATC Forward primer to amplify PtEG16 promoter with corrections shown by sequencing CCTGCAGGTCGACTCTAG Forward sequencing primer for pMDC162 GTTGGGGTTTCTACAGGACG Reverse sequencing primer for pMDC162 from GUS gene CAAGGCGATTAAGTTGGG Forward sequencing primer for pMDC162 181 GATCAATTCCACAGTTTTCGC Reverse sequencing primer for pMDC162 from GUS gene AATTGTTGGTCTTCAACGAGGAA Forward primer for amplifying housekeeping gene 18S AAAGGGCAGGGACGTAGTCAA Reverse primer for amplifying housekeeping gene 18S GACAAATCCCCAAGACCATCG Forward primer to amplify part of EG16 RNAi fragment bridging exons for mRNA quantification GAATCACAAGAACACTCCACCG Reverse primer to amplify part of EG16 RNAi fragment from end of gene CCACAGTTTTCGCGATCCAG Genotyping RC19, anneals to beginning of GUS gene CGCGGTTAGCACAAACAAAC Genotyping RC19, anneals to end of promoter CGCTGCAGTTATCATCATC Genotyping RC8 GTGAGGAAAGTGGAGAAAAGGG Genotyping RC8, anneals to EG16 RNAi fragment sequence 182 Figure D.1. EG16 expression in Populus trichocarpa tissues. (A). Microarray expression analysis from the eFP browser (Winter et al., 2007). (B). RNA-seq expression data from the PopGenIE database (Sjödin et al., 2009, Sundell et al., 2015). 183 Figure D.2. Genotyping of hybrid poplar P39 transformants. (A). Amplification of part of the PtEG16 promoter and GUS gene sequence from gDNA extracted from P39 pPtEG16::GUS lines. The expected size is 567 bp. (B). Amplification of the RNAi insertion from gDNA extracted from EG16-downregulated (RC8) P39 lines. The expected size is 441 bp. 184 Figure D.3. Western blot for anti-PtEG16 antibodies. Western blot of serially diluted recombinant PtEG16 purified protein produced in E. coli (1 – 0.001 µg) using anti-PtEG16(full-length) (Cedar Lane, Burlington, Ontario, Canada) (A) and anti-PtEG16(peptide) antibody (Agrisera, Vännäs, Sweden) (B). 185 Figure D.4. Immunofluorescence microscopy using anti-PtEG16(full-length) antibody. Sectioned petiole tissue from greenhouse-grown plants was used in whole-mount fluorescence microscopy. Young petioles were collected from the 2nd node and mature petiole from the 8th node. As Solanum lycopersicum (tomato) tissues do not have an EG16 homolog, they served as controls. Scale bars represent 2 mm. 186 Figure D.5. Design of PtEG16 peptide used as the antigen to produce the anti-PtEG16(peptide) antibody. (A). Alignment of PtEG16 protein sequence to Populus trichocarpa XTH sequences. Purple highlights show the PtEG16 peptide sequence and matching amino acids. (B). Phyre 2.0 model of PtEG16 highlighting the peptide sequence location in purple. 187 Figure D.6. RT-qPCR of wild-type and EG16 RNAi lines. Error bars indicate the standard deviation of three technical replicates. a – not significant, b – statistical significance as calculated by a one-tailed Student’s T-Test. 188 Figure D.7. Assessment of MLG in P39 poplar tissues. (A). Confocal microscopy of P39 poplar sections immunostained with anti-MLG (BG1) antibody. Scale bars represent 100 µm (B). MALDI-imaging spectra of P39 poplar wild-type and RNAi. 189 Figure D.8. MALDI-imaging spectra of xyloglucan oligosaccharides in wild-type (A) and RNAi-EG16 P39 hybrid poplar (B). 190 Figure D.9. Soluble sugar analysis of leaves from 3 months old greenhouse plants. Young leaves refer to leaves from 1st to 5th node, and mature is 8th node. Each bar represents three biological replicates and technical duplicates.
Thesis/Dissertation
eng
Biochemistry and Molecular Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-ShareAlike 4.0 International
Graduate
Brumer, Harry
Understanding the diversity, biochemistry, and biological role of endo-glucanase 16 enzymes in the context of plant evolution
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