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Carbon dioxide enrichment and the role of carbohydrate reserves in root growth potential of cold-stored… Chomba, Bernard Malata 1992

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CARBON DIOXIDE ENRICHMENT AND THE ROLE OF CARBOHYDRATE RESERVES IN ROOT GROWTH POTENTIAL OF COLD-STORED ENGELMANN SPRUCE (PICEA ENGELMANNII PARRY) SEEDLINGS  by Bernard Malata Chomba B.Sc.(For.) (Hons.) University of Dar-es-Salaam, Tanzania  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE  in THE FACULTY OF GRADUATE STUDIES (DEPARTMENT OF FOREST SCIENCES)  We accept this thesis as conforming to the required standard  THE UNIVERSITY OF BRITISH COLUMBIA April 1992 © Bernard Malata Chomba, 1992  In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission.  Department of  ^  ct^cF- s  The University of British Columbia Vancouver, Canada Date  DE-6 (2/88)  AT^CY)-  ii  ABSTRACT Two experiments were conducted to examine the role of carbohydrate reserves in spring root growth potential (RGP) of Engelmann spruce (Picea engelmannii Parry) seedlings. In the first experiment, the effects of pre-storage carbon dioxide enrichment (CE) on total non-structural carbohydrates (TNC) and post-storage root growth were studied. Seedlings were grown from seed for 202 days in growth chambers with ambient (340 gL•L -1 ) and CO2 enriched (1000 gL•L -1 ) air. Reciprocal transfers between treatments took place at 60 and 120 days. Photoperiod was reduced at 100 days to induce bud set. After 180 days seedlings were hardened-off for storage at -5 ° C. At 268 and 327 days, seedlings were planted in a growth chamber in three water baths. New roots >5 mm long were counted after 28 days growth. Seedlings were also assessed for bud break every two days. At each planting time, and at 80, 120, 140, and 202 days, seedlings were randomly selected from each of the CO2 treatments and harvested for analysis of starch and soluble sugar content. Growth data were also collected. In the second experiment, the relative contributions of reserve carbon and current photosynthate to new root growth were studied. Seedlings were raised using standard nursery procedures up to bud set (end of September, 1990). Seedlings were then moved into a growth chamber and placed in four Plexiglas boxes for stable carbon isotope labelling. Two boxes received ambient CO2 with normal isotopic composition  iii  and the other two received air that was first stripped of CO 2 before adding back tank CO2 depleted in 13 C. On November 30, seedlings were hardened-off for storage for four months at -5 °C. Samples of seedlings were taken both before and after storage and analyzed for carbohydrate reserves. After storage, seedlings were planted in a growth chamber for 36 days, during which time seedlings were sampled for new root growth at 9, 18, and 36 days. Extracted carbohydrates and new white roots were then analyzed for 13 C/ 12 C ratios. Carbon dioxide enrichment increased seedling biomass and root collar diameter, more so after bud set. Stem height was only slightly affected by CE. Shoot:root ratios (DW basis) were not affected by CE, but decreased steadily with age. Carbon dioxide enrichment had little effect on soluble sugars, starch, and TNC prior to bud set. After bud set, CE greatly promoted the rate of reserve accumulation, but did not influence the final level attained prior to storage. Needles were the major storage organ for soluble sugars, while roots were so for starch. Soluble sugars were not strongly affected by the two and four months storage. In contrast, there was more than 50% reduction in starch in needles, stems, and roots, and almost a one-third loss of TNC during storage. None of the CO2 treatments had any influence on bud break and RGP. The isotope labelling experiment indicated that carbohydrate reserves did make an important contribution to spring root growth, but this sink was only a minor drain on the reserve pool.  iv  TABLE OF CONTENTS Page ABSTRACT ^ Table of Contents ^  iv  List of Tables ^  vi  List of Figures ^  vii  List of Appendices ^  ix  Abbreviations ^ Acknowledgements ^  xi  Dedication ^  xii  1.  INTRODUCTION ^  1  2.  LITERATURE REVIEW ^  4  2.1. Carbohydrate Reserves and Root Growth ^ 4 2.2. Carbohydrate Reserves and CO2 Enrichment ^ 8 2.3. Carbohydrate Reserves and Root Growth Potential (RGP) During Storage ^ 10 2.4. Stable Carbon Isotopes as Tracers ^ 12 2.5. Soil Temperature and Root Growth ^ 13 15 2.6. Objectives ^ 3. MATERIALS AND METHODS ^  16  3.1. Experiment One: Carbon Dioxide Enrichment 16 (CE) and Carbohydrates ^ 3.1.1. Seedling Production and Cold-storage ^ 16 3.1.2. Post Cold-storage Planting ^ 19 3.1.3. Carbohydrate Analyses ^ 22 3.1.3.1. Soluble Sugars Analysis by Anthrone Reagent ^ 22 3.1.3.2. Starch Analysis by Enzymatic 23 Hydrolysis ^ 3.2. Experiment Two: Stable Carbon Isotopes 24 as Tracers ^ 3.2.1. Seedling Production and Cold-storage ^ 24 3.2.2. Post Cold-storage Planting ^ 27 3.2.3. Stable Carbon Isotope Analyses ^ 29 3.2.3.1. New White Roots ^ 31 3.2.3.2. Starch Extraction ^ 32 3.2.3.3. Soluble Sugars Extraction ^ 34 3.3. Experimental Design and Data Analysis ^ 35 3.3.1. Experiment One ^ 35  3.3.2. Experiment Two ^ 4.  RESULTS ^  37 38  4.1. Carbon Dioxide Enrichment (CE) Experiment ^ 38 4.1.1. Effects of CE on Growth Prior to 38 Storage ^ 4.1.1.1. Biomass Accretion and Partitioning ^ 38 4.1.1.2. Root Collar Diameter and Height Growth ^ 44 4.1.2. Carbohydrate Reserves ^ 46 4.1.3. Bud break and Root Growth Potential ^ 57 4.2. Stable Carbon Isotopes as Tracers ^ 60 5.  DISCUSSION ^  65  Biomass Accretion and Partitioning ^ 65 Root Collar Diameter and Stem Height ^ 67 Carbohydrate Reserves ^ 68 Effects of Cold-storage on Carbohydrate Reserves ^ 72 5.5. Bud Break ^ 77 5.6. Root Growth Potential (RGP) ^ 80 5.6.1. Effects of CE and Soil Temperature on RGP ^ 80 5.6.2. Effects of Cold-storage on RGP ^ 82 5.6.3. Role of Carbohydrates in RGP ^ 84  5.1. 5.2. 5.3. 5.4.  6.  CONCLUSIONS ^  88  7.  RECOMMENDATIONS ^  90  8. LITERATURE CITED ^ APPENDICES ^  93 103  vi  LIST OF TABLES Page  Table^  1.  The effects of CO2 enrichment (CE) on days to first bud break in Engelmann spruce seedlings ^ 58  2.  The effects of CE on post-storage root growth potential (number of new roots > 5 mm long) in Engelmann spruce seedlings ^  3.  59  Isotopic composition (expressed as 8 13 C value) of whole seedling total tissue and TNC before and after storage ^  4.  61  Trends in isotopic composition (expressed as 8 13 C value) of new white roots, and calculated per cent of reserve carbon to new root construction of "labelled"  seedlings  62  vii  LIST OF FIGURES Figure^  Page  1.  A general view of the CE experiment during the seedling production phase ^  17  2.  A general view of the pot arrangement for the CE experiment in the growth chamber in one of the water baths ^  21  3.  A schematic diagram of the equipment used in the stable carbon isotope labelling experiment ^ 25  4.  General views of the arrangement when seedlings were moved into the Plexiglas boxes for isotope labelling (a), and during post-storage planting (b) ^ 28  5.  Effects of CE on (a) needle dry weight, and (b) stem dry weight of Engelmann spruce seedlings ^ 40  6.  Effects of CE on (a) root dry weight, and (b) total seedling biomass of Engelmann spruce seedlings ^  42  7.  Effects of CE on shoot:root ratio (DW basis) of Engelmann spruce seedlings ^  43  8.  Effects of CE on seedling mean diameter (a), and stem height (b) of Engelmann spruce seedlings ^ 45  9.  Trends in whole plant TNC content of Engelmann spruce seedlings ^  47  10. Trends in allocation of reserves to individual plant parts expressed as a percentage of the maximum TNC observed prior to storage ^ 48 11. Trends in soluble sugar content of needles (plus buds) of Engelmann spruce seedlings ^ 50 12. Trends in the soluble sugar content of stems of Engelmann spruce seedlings ^  51  13. Trends in soluble sugar content of roots of Engelmann spruce seedlings ^  52  14. Trends in starch content of needles (plus buds) of Engelmann spruce seedlings ^  54  15. Trends in starch content of stems of Engelmann spruce seedlings ^  55  viii 16.  Trends in starch content of roots of Engelmann spruce seedlings ^  17.  Glucose (a), and starch (b) standard curves used in carbohydrate analyses ^  56 120  ix  LIST OF APPENDICES Appendix^  Page  1.  ANOVA for the effect of CE on biomass partitioning in Engelmann spruce seedlings ^  2.  ANOVA for the effect of CE on total seedling biomass production in Engelmann spruce seedlings ^ 105  3.  ANOVA for the effect of CE on shoot:root ratio in Engelmann spruce seedlings ^  4.  ANOVA for the effect of CE on diameter and stem height in Engelmann spruce seedlings ^ 107  5.  ANOVA for the effect of CE on carbohydrate reserves in Engelmann spruce seedlings at 80 days ^ 108  6.  ANOVA for the effect of CE on carbohydrate reserves in Engelmann spruce seedlings at 120 days ^ 109  7.  ANOVA for the effect of CE on carbohydrate reserves in Engelmann spruce seedlings at 140 days ^ 110  8.  ANOVA for the effect of CE on carbohydrate reserves in Engelmann spruce seedlings at 202 days ^ 111  9.  ANOVA for the effect of CE on carbohydrate reserves in Engelmann spruce seedlings at 268 days ^ 112  103  106  10. ANOVA for the effect of CE on carbohydrate reserves in Engelmann spruce seedlings at 327 days ^ 114 11. ANOVA for the effect of CE on bud break and RGP in Engelmann spruce seedlings at 268 and 327 days....115 12. Protocol for carbohydrate analyses ^  116  LIST OF ABBREVIATIONS ANOVA^  Analysis of variance  cc^  Cubic centimetre  CE^  Carbon dioxide enrichment  DF^  Degrees of freedom  DW^  Dry weight  g  Gravitational acceleration  i.d.^  Inside diameter  MOF^  Ministry of Forests  MS^  Mean square  o.d.^  Outside diameter  P  Probability  RGP^  Root growth potential  SE^  Standard error of the mean  SS^  Sum of squares  SV^  Source of variation  TNC^  Total non-structural carbohydrates  $'0^  Per mil or per thousand  813c^  Stable isotope abundance parameter  xi  ACKNOWLEDGEMENTS I wish to express my profound gratitude to my supervisor, Dr. R.D. Guy for the academic guidance rendered throughout the course of this study. His academic enthusiasm, tireless guidance, and patience will always be remembered. I am equally deeply indebted to the other members of my supervisory committee, Dr.'s D.P. Lavender, and P.A. Jolliffe who did not only review this manuscript, but gave both material and moral support to bring this study to this stage. Special thanks go to Dr. H.G. Weger for his diligent advice and guidance on carbohydrate analyses, and Dr. S.N Silim for advice and assistance on various laboratory techniques and procedures. I am also grateful to all the staff, graduate students and workers of Ponderosa Annex B for all the co-operation extended to me in various ways. I am particularly grateful to A. Balisky who so kindly proof read this manuscript, and from time to time came to my rescue whenever I ran into computing problems. I wish to thank the International Development Research Centre (IDRC, Canada) in co-operation with the Zambian Government for the financial support that made this study possible. I am also grateful to Pacific Regeneration Technologies Inc. who funded a major part of my thesis research. I would also like to thank the Zambian community in Vancouver and the African students at UBC for the kindness, understanding, and co-operation that I shared with them. Finally, I wish to extend my sincere thanks to my wife, Cecilia, and my two sons, Ian Musakanya and Chama Kizito for their patience, perseverance, and encouragement during my academic struggle. I miss them a million times.  xii  This thesis is dedicated to my second born son, CHAMA KIZITO CHOMBA whom I left when he most needed me.  1 1.0 INTRODUCTION.  Rapid initiation of adequate and vigorous root growth is necessary for early survival and establishment of outplanted seedlings (Burdett 1987, Burr 1989, Ritchie et al. 1985, Sutton 1987). Root initiation and early growth will depend on environmental factors (e.g. soil moisture, soil temperature, nutrients, photoperiod, etc.), the seedlings' physiological status (e.g. cold hardiness, bud dormancy, stress resistance, carbohydrate reserves, etc.), and management practices (e.g. lifting date, freezer-storage duration, site preparation, etc.). This study attempts only to examine the role of carbohydrate reserves in spring root growth of freezer-stored Engelmann spruce (Picea engelmannii Parry) seedlings. Emphasis is placed on investigating the utility and possible side effects of carbon dioxide enrichment (CE) on the manipulation of these reserves. During the past twenty years, the reforestation program in British Columbia has been characterized by the planting of nursery produced seedlings. Prior to 1975, most planting took place in the coastal areas of British Columbia. More recently, planting efforts in the interior of the province have become more important and the species currently predominant in the reforestation program include white spruce (Picea glauca (Moench.) Voss), Engelmann spruce, and lodgepole pine (Pinus contorta Loud.). Furthermore, the majority of the seedlings planted in the interior are, in fact, produced at the southern latitudes and lower  2  elevations. For example, during the 1987 planting season, interior spruce (white spruce and Engelmann spruce, and are hybrids) and lodgepole pine together made up well over 75 per cent of the approximately 230 million container seedlings planted in British Columbia (B.C. MOF 1989, Lavender unpublished data). For the 1990 planting season only 208 million seedlings were planted, and the proportion of interior spruce was comparable to the figures of the previous years. This shift in the emphasis of the reforestation program has necessitated the development of cold/frozen storage of fall or early-winter lifted seedlings. In cold-storage, the seedling's dormancy and resistance to stress may be maintained until planting time in May/June. If the seedlings are not kept cold, they will break their buds in spring in response to the rising temperature, before the snow melts in the interior. The current practice is to cold/freeze store interior spruce seedlings four to eight months, prior to outplanting them in spring or early summer. Lack of root growth by outplanted interior spruce seedlings has been noted as one of the major factors contributing to plantation failure in north-central British Columbia, where several hundred thousands of hectares are not satisfactorily restocked (Butt 1986). This lack of root development may have a direct bearing on the reduction of the ability of outplanted seedlings to take up necessary moisture from the soil, and is presumed to cause "planting  3  check" (Grossnickle 1988). Although root growth may be correlated with high survival and growth of outplanted seedlings (Burdett 1979, Stone 1955), there seems to be no adequate data to support and explain how the cause-effect relationship is mediated (Binder et al. 1988, Lavender 1988). Initial survival and early growth of outplanted seedlings may largely depend on the physiological readiness of the seedlings to rapidly regenerate new roots in order to establish intimate contact with the soil matrix, thereby being able to resume water and nutrient uptake. The ability of seedlings to promptly and abundantly initiate and/or elongate new roots after outplanting in a favourable environment has been referred to by many authors as root growth potential (RGP) or root growth capacity (RGC) (Cleary et a/. 1978, Marshall 1985, Ritchie 1982, 1984, Stone 1955). However, the relationships between RGP and the seedling's field performance have not been adequately researched (Binder et al. 1988, Lavender 1988, Ritchie and Tanaka 1990), and also not all tests show positive relationships (Binder et a/. 1988, Landis and Skakel 1988). It is therefore important to note that RGP is only a measure or good predictor of overall seedling vigour (Lavender 1988, Ritchie and Dunlop 1980, Stone 1955). As such, any interpretation of RGP test results for predicting outplanting field performance must consider other factors such as site conditions and planting quality (Binder et a/.  4  1988). Furthermore, RGP is not a physiological process per  se; it is an integrated manifestation of many important physiological processes (e.g. cold hardiness, stress resistance, carbohydrate status, etc.). For this reason RGP has become a popular and useful indicator of seedling vigor and stress resistance; the rationale being that, if there is any problem with seedling physiology, it should show up as a decrease in the seedling's ability to produce new roots (Lavender 1988, Ritchie 1987).  2.0 LITERATURE REVIEW 2.1 Carbohydrate Reserves and Root Growth Root growth can only proceed at the expense of available metabolic substrates, primarily carbohydrates. In the case of conifers, lipids and proteins are also important reserve materials for root growth. Carbohydrate reserves play an important role in maintaining respiration and growth during times when photosynthesis is not taking place (Duryea and McClain 1984, Kramer and Kozlowski 1979, Ritchie 1984). According to Ritchie and Dunlop (1980) and Duryea and McClain (1984), the most abundant translocatable carbohydrate in trees is sucrose. On the other hand, starch  is the most abundant storage form of carbohydrate. Furthermore, plants are capable of accumulating and storing carbohydrates in roots, stems, and foliage for subsequent metabolism either within the site of assimilation and storage or for export to other growing tissues (Glerum  5  1980a, Kramer and Kozlowski 1979). In terms of root development, it is not well known whether root growth at planting time proceeds at the expense of stored carbohydrates, at the expense of recently photoassimilated carbohydrates, or both. Several studies on the role of carbohydrate reserves in root growth potential have presented conflicting results. For example, Ritchie and Dunlop (1980) concluded that initiation of new roots in most conifer seedlings depends on substances (presumably current photosynthate and growth hormones) originating in the shoots and translocated through the phloem to the root system. In addition, many studies on root development in Douglas-fir (Pseudotsuga menziesii (Mirb.) Franco) have shown that the main source of stimulus for spring root growth is the foliage (Philipson 1988, Ritchie 1982, van den Driessche 1987). Results from bark-ringing experiments by Zaerr and Lavender (1974) indicated that food reserves in roots of the girdled Douglas-fir seedlings declined steadily during the five weeks of the experiment but root activity dropped to zero within three weeks. Results from the above experiment led Zaerr and Lavender (1974) to conclude that root activity may require carbohydrates but the level of food reserves alone does not control root growth. Similarly, results from radioisotope experiments with Douglas-fir and Sitka spruce (Picea sitchensis (Bong.) Carr), and low CO2 concentration and girdling experiments with Douglas-fir seedlings, led van den Driessche (1987, 1991) to speculate  6  that current photosynthate is the primary carbon source for early root growth. Other researchers have found that phytohormones may also significantly influence early root growth. For example, Kramer and Kozlowski (1979) stated that, as dormancy intensity weakens in spring, buds export increasing amounts of auxins and gibberellins, which in conifers may move to leaves and stimulate the production or accumulation of root promoters. In deciduous species where no leaves are present, rooting promoters remain in the buds or stems (Carlson and Larson 1977). In addition, in barkringing experiments, Zaerr and Lavender (unpublished data) demonstrated that some substance (most likely plant growth regulators) besides current photosynthate controlled root growth because  14 C-sucrose  was fed to the lower edge of the  girdle, roots did not grow, but radioactivity was isolated in the root tips. In both conifers and hardwoods, growth hormones move downward through the phloem parenchyma to initiate root growth (Salisbury and Ross 1985). Work on ponderosa pine (Zaerr 1967), Douglas-fir (Deyoe and Zaerr 1976), and red oak (Quercus rubra L.) (Carlson and Larson 1977) seedlings also confirmed that plant growth regulators (particularly auxins and gibberellins) enhanced root initiation. In nature, mycorrhizae are also important to root growth of higher plants by way of increasing the efficiency of nutrient and water uptake, and may affect plant growth hormone physiology (Harley and Smith 1983, Mikola 1980).  7  On the other hand, other researchers claim that new root growth largely depends on carbohydrate reserves. For instance, Ronco (1973) found that first year survival of outplanted Engelmann spruce seedlings seemed to depend on food reserves accumulated while in the nursery. In addition, Kozlowski and Keller (1966), and Krueger (1967) speculated that since the rate of carbon assimilation of newly planted seedlings is rather low until the time they become established, it is logical to assume that new root growth depends on stored carbohydrates. Marshall (1985) expressed the view that seedlings are dependent on reserve carbohydrates from the time they are lifted until photosynthesis is sufficient to meet the demands of growth and respiration. He further pointed out that if carbohydrate reserves are inadequate to meet the respiratory demands associated with the cold storing and outplanting of the seedlings, the seedlings will die. In a bark-ringing experiment by Philipson (1988), root growth in Sitka spruce seedlings showed dependance on carbohydrate reserves, while root growth in Douglas-fir seedlings depended entirely on recently assimilated photosynthate. Contrary to the above for Douglas-fir, findings in support of carbohydrate reserves being responsible for early root growth following storage were reported by Krueger and Trappe (1967). Based on the available literature, it appears that the results obtained so far have mostly been with coastal species, which, in terms of overwintering strategy, may be quite  8  different from interior species.  2.2 Carbohydrate Reserves and CO2 Enrichment  One way to assess the role of carbohydrate reserves in RGP is by producing seedlings with different levels of starch reserves prior to placing them in cold-storage. One possible way to increase carbohydrate reserves is through CO 2 enrichment of the seedlings before taking them into cold-storage. The positive effects of short-term CO2 enrichment on the growth and yield of plants have been widely demonstrated (Campagna and Margolis 1989, Gifford 1979, Jolliffe and Ehret 1984, Mortensen 1987, and Waggoner 1984). According to Mortensen (1987), the optimal CO2 concentration for plant growth and yield ranges from 700 to 900 gL•L -1 CO 2 . This is approximately two to three times higher than the current atmospheric CO2 concentration (ca. 340 gL•L -1 CO2). However, the responses of plants to increasing CO2 concentration will mostly depend on environmental factors such as soil moisture, nutrients (particularly nitrogen and phosphorus), soil temperature, and light intensity. For example, Brown and Higginbotham (1986) found that CO2 enrichment significantly increased total dry weight of white spruce and aspen (Populus tremuloides Michx) seedlings grown in a high nitrogen regime as opposed to a low nitrogen regime. Conroy et al. (1990) also indicated that, both high phosphorus and CO 2 levels increased the total dry weights  9  and growth rates of Pinus taeda D. Don and Pinus caribaea var. hondurensis seedlings, and that these effects were synergistic. Other researchers found positive and synergistic effects of CO 2 enrichment and high irradiance on the growth rates and biomass production of seedlings (Tolley and Strain 1984, Yeatman 1970). For instance, Tolley and Strain (1984) reported that elevated CO 2 concentration (ca. 650 gL•L -1 CO 2 ) and high irradiance (ca. 1000 gmol quanta m-2s-1 ) increased total dry weight of sweetgum (Liquidambar styraciflua L.) and Pinus taeda L. seedlings. In both  species, these effects were highly dependent on age (the younger the seedlings the more responsive they were), and duration of exposure (prolonged exposure resulted in no treatment effects). It is also important to note that the effects of elevated CO2 concentration are species specific, with some species responding more positively than others (Brown and Higginbotham 1986, Kramer 1981, Sionit et al. 1985, Tolley and Strain 1984). Results from CE experiments with black spruce (Picea mariana Mill.) by Campagna and Margolis (1989) showed no  significant differences in the concentrations of sugars, starch, or total nonstructural carbohydrates (TNC) in either roots or stems after three to six weeks exposure to elevated CO2 levels. On the other hand, CE significantly increased starch concentrations in the needles. Furthermore, Yeatman (1970) demonstrated that elevated CO2 concentrations with high irradiance increased the seedling dry weight of species  10  such as white spruce, Norway spruce (Picea abies (L.) Karst), jack pine (Pinus banksiana Lamb.), and Scots pine (Pinus sylvestris L.), by 30 to 80 per cent at 1,000 µL•L -1  CO2. Campagna and Margolis (1989), and Surano et al. (1986) also found similar responses with black spruce and ponderosa pine (Pinus ponderosa Dougl. ex P. Laws.) seedlings respectively. However, very little is known about the effect of CO 2 enrichment on starch concentration in conifer seedlings. The optimum stage at which CO2 enrichment will effectively increase carbohydrate reserves during seedling production in the nursery is also not known. For example, virtually none of the early work on CE has examined the effects of CE on carbohydrate reserves after bud set, when height growth ceases but photosynthesis continues. Moreover, according to the views of many researchers (Hollinger 1987, Kramer 1981, Surano et al. 1986), in forestry, the long term effect of elevated CO2 concentrations on the growth and physiological conditions of seedlings has received too little attention. The present study attempts to examine the effects of CE on carbohydrate reserves prior to coldstorage, and on post-storage RGP in Engelmann spruce seedlings.  2.3 Carbohydrate Reserves and RGP During Storage Duration of cold-storage has been implicated in the appearance of adverse effects on the physiological condition of seedlings. For instance, Loescher et a/. (1990),  11  McCracken (1979), Ritchie (1982), and van den Driessche (1979) stated that one important change occurring in coldstored seedlings, which might account for changes in performance, is the gradual respiratory depletion of reserve sugar and starch. This stems from the fact that stored seedlings,^which are no longer able to actively photosynthesize,^slowly metabolize their carbohydrate reserves. Loss of such reserves has been implicated as being responsible for poor survival and poor RGP (Hellmers 1962, Stone 1955, 1970, Stone and Jenkinsen 1970). Carbohydrate depletion in stored seedlings has been extensively studied in Douglas-fir by Ritchie (1982), in jack pine by Glerum (1980b), in ponderosa pine by Hellmers (1962), in Engelmann spruce by Ronco (1972, 1973), and in mugo pine (Pinus mugo Turra var. mughus) and in radiata pine (Pinus radiata D. Don) by McCracken (1979). All these authors hypothesized that poor seedling performance at planting time was correlated with a depletion of carbohydrate reserves during cold-storage. For interior spruce seedlings, Ritchie (1985, 1987) and Ritchie et a/. (1985), indicated that RGP is related to dormancy release. Generally speaking, RGP increases as buds accumulate chilling hours and finally peaks when the chilling requirement has been fulfilled. Thereafter, it declines rapidly, as the internal metabolic focus of the seedlings switches to the elongation of the new shoots (Ritchie and Dunlop 1980, Ritchie and Tanaka 1990).  12  2.4 Stable Carbon Isotopes as Tracers  Another approach to resolving the carbohydrate reserve controversy and establishing the absolute contributions of old and new photosynthate to post-storage root growth, is by the use of stable isotopes. Until recently, most C-labelling experiments have employed radioactive 14 C (van den Driessche 1987, Glerum 1980a, Ursino et a1. 1968), but regulatory constraints associated with use of radioactive materials make some such studies awkward, particularly long-term labelling experiments and field studies (Svejcar et al. 1990). In addition to regulatory constraints, the period for radioactive 14 C labelling is short which may result in insufficient or uneven labelling of plant material. On the other hand, because of the advantages of safety, longer period of labelling, and increased availability of isotope ratio mass spectrometers, stable carbon isotope techniques are beginning to gain in popularity for use in tracer studies in plants (Svejcar et al. 1990, Mordacq et a/. 1986, O'Leary 1981). As used in this study, the technique involves growing seedlings, prior to storage, in air containing CO2 with different proportions of 13 C and 12 C. Later, the changes in the 13 C/ 12 C ratio can be determined in the new  roots produced after the storage period (Glerum 1980a, Svejcar et al. 1990). If after cold-storage such seedlings are allowed to grow under normal air, all new photosynthate will contain a different 13 C/ 12 C ratio. Stable isotope labelling approaches such as this, particularly at the  13  natural abundance level, have not been extensively used in studies of plant/tree physiology and the present study is somewhat unique in trying to explore these possibilities.  2.5 Soil Temperature and Root Growth  Regardless of the condition of the seedlings, soil temperature may exert a major influence on root development and ultimately establishment. Studies by Heninger and White (1974), and Hermann (1962) revealed that expression of RGP in transplanted seedlings was highly dependent on soil temperature. Camm and Harper (1991) also found that root growth of over-wintered, cold-stored white spruce seedlings was temperature dependent. However, it is interesting to note that the effect of soil temperature on root development appears to be species specific such that different species have different optimal soil temperatures. As noted by Ritchie and Dunlop (1980), and Lavender and Overton (1972), root development proceeds favourably between 18 and 25 ° C, depending upon the species, with those species native to cooler climates tending to have a lower optimal soil temperature (e.g. interior spruce: 19 ° C) than those native to warmer climates (e.g. loblolly pine: 25 ° C). On the other hand, Husted and Lavender (1989) reported relatively lower optimal soil temperature for white spruce (e.g. 15-17 ° C) than the figures quoted above for interior spruce. In the interior, soil temperatures at planting time are generally far below the above mentioned figures. The effect  14  of such low soil temperatures on plantation establishment is through a slowing of early root development and shoot growth (DeLucia 1986, Grossnickle 1988, Grossnickle and Blake 1985), presumably due to decreased metabolic activity and a decreased turgor of root cells caused by reduced water uptake (Lopushinsky and Kaufmann 1984). According to Lopushinsky and Kaufmann (1984), the main causes of decreased uptake of water at low soil temperatures are increased viscosity of water and decreased permeability of root cell membranes. Furthermore, there are speculations that carbohydrate reserves play an important role in osmotic adjustment for purposes of increasing water uptake in conifer roots growing at low temperatures (Ritchie 1982, DeLucia 1986). The occurrence of high carbohydrate levels in roots of seedlings grown at low soil temperature as opposed to high soil temperature, may in part, reflect this role. For example, Weger and Guy (1991) demonstrated that both soluble sugar and starch levels in white spruce were approximately 50% higher in 4 ° C-grown roots than 11 and 18 ° C-grown roots. Marshall and Waring (1985) reported similar trends in starch and soluble sugar content in the roots of Douglas-fir seedlings grown at 10, 20, or 30 ° C soil temperatures. On the other hand, experiments with Douglasfir (Ritchie 1982) and Engelmann spruce (DeLucia 1986) seedlings revealed that conversion of starch to sugars in roots is an important acclimation response to low temperatures. The rationale being that, sugars depress the  15  freezing point of the cell sap, tend to concentrate in the vacuole and reduce the probability of intracellular ice crystal formation, and may serve some cryoprotective role with respect to cell membranes (Santarius 1982). For this reason, it is probably right to assume that carbohydrates play an important role in frost hardiness. As noted by many authors (Guy 1989, Levitt 1980, Santarius 1982), accumulation of free sugars in both woody and perennial plants seems to be a requirement for cold acclimation.  2.6 OBJECTIVES  Based on the foregoing literature review, this study was conducted with the following objectives in mind: 1. To investigate the effects of CE on carbohydrate levels and post-storage root growth in Engelmann spruce seedlings. In addition, the effects of CE on other morphological and physiological characters were examined. 2. To observe changes in carbohydrate levels during coldstorage. 3. To measure relative proportions of old carbon (from stored carbohydrates) and new carbon (from current photosynthate) in new roots of cold-stored Engelmann spruce seedlings.  16  3.0 MATERIALS AND METHODS  The study consisted of two experiments. The first experiment involved producing seedlings under six CO2 regimes and then storing them during winter for two different storage durations. This was followed by planting the seedlings in growth chambers at 268 and 327 days (i.e. after two and four months of storage, respectively). The second experiment involved producing seedlings in the nursery using the standard nursery practices up to bud set (September). Thereafter, seedlings were differentially labelled with 13 CO2 until the seedlings were ready for coldstorage.  3.1 Experiment One: CO 2 Enrichment and Carbohydrates 3.1.1 Seedling Production and Cold-storage  Engelmann spruce seedlot number 8356^(location: Spapilum, 51 ° 14' latitude, 119 ° 39' longitude) was sown in May, 1989 in the Plant Science Laboratory at UBC. The seeds were sown in eight home-made growth cabinets belonging to Dr. Jolliffe, Plant Science Department, and CO2 enrichment commenced immediately. The CO 2 regimes consisted of high and low concentrations (i.e. 1000 gli•L -1 and 340 1.1L•L -1 of CO2 respectively), and reciprocal transfers of seedlings from low to high and high to low CO2 concentrations. Transfers were conducted after 60 and 120 days of CO2 treatment. Air was pulled into the chamber from outside the MacMillan building by fans mounted in the base of each cabinet (Fig.  17  1). Two chambers were used and each one of them was divided into 4 compartments, with replicate treatments arranged  Figure 1. A general view of the CE experiment during the seedling  production phase.  18  diagonally opposite each other. Tank CO2, regulated manually by adjusting rotometer valves, was added to the incoming air for the high CO 2 cabinets. Carbon dioxide concentrations in all the cabinets were monitored with an infrared gas analyzer (IRGA, LI-864 CO2 Analyzer from Beckman, Fullerton, CA). Air temperature and humidity in each chamber were recorded continuously with a Campbell Scientific (Logan UT) 21X data logger. On average the growth cabinet environment was as follows: day and night temperatures 22/15 ° C, 18 hour photoperiod initially and then reduced to 9 hours at 100 days to induce bud set, which followed within 2-3 weeks, photosynthetic photon flux density ca. 190 Rmol quanta m -2 s 1 (full sunlight is about 2000 Rmol quanta m -2 s -1 ), and relative humidity 70-75% during the day and 85-90% at night. Initially, the root medium contained slow release fertilizer, but seedlings were also fertilized on a regular basis with 20:8:20 N:P:K and micronutrients. Seedlings were grown for 180 days following emergence, then hardened off. Seedlings were hardened off by moving them into water-cooled Plexiglas fumigation boxes within a Conviron E15 (Winnipeg, Manitoba) growth chamber. Day/night temperatures were reduced to 8/5 ° C. Other conditions remained the same and the CO2 treatments were continued for three weeks, by which time cold hardiness had reached at least -22 ° C (Silim: personal communication). Thereafter, seedlings were packed and then taken into cold-storage (at -5 ° C) for two and four months. The first transfer was done at 60 days and the second  19  one at 120 days. At 80, 120, 140, and 202 days, some seedlings were randomly sampled from each treatment. The sampled seedlings were killed in liquid nitrogen, freezedried, and then they were kept in a freezer for later carbohydrate analysis. At each harvest date, growth data were also collected. At 202 days, the remaining seedlings were sorted according to treatment, packed and taken into cold-storage (-5 ° C). There were other harvests prior to 80 days, but these are not included in the thesis because emphasis was placed on the examination of CE effects after transfers (i.e. 60 and 120 days) and bud set.  3.1.2. Post Cold-storage Planting  At 268 and 327 days, seedlings were removed from coldstorage and allowed to thaw for seven days (at 5 ° C). Seedlings were then randomly selected, potted, and placed in three water baths (3 ° C, 7 ° C, 11 ° C for 268 days; and all at 11 ° C for 327 days) in a Conviron E30 (Winnipeg, Manitoba) growth chamber at the UBC South Campus Nursery. The intention was to have another planting at 388 days (i.e. after 6 months storage), but the seedlings scheduled for this planting date were spoiled when the cold-room failed. For the root medium, peat/perlite (2:1) was used with dolomite lime added to the mixture to adjust the pH. To raise the root medium pH of approximately 4.65 to about 5.5, 600 g of dolomite lime was added to 20 L of peat/perlite mixture. Silica sand (800 mL) was placed at the bottom of  20  each pot to keep the pots in place. There were nine pots per water bath (all six CO2 treatments represented) with four seedlings per pot (Fig. 2). Styrofoam covers were installed over each water bath to minimize heat transfer to and from the air around pots. The styrofoam covers also facilitated keeping the pots in place. A thermometer was placed in each water bath to monitor the root zone temperature. Another thermometer was suspended above the seedlings in each water bath to cross-check the room air temperature with the temperature set on the growth chamber control system. No fertilizer was added to either the root medium or the water, because 28 days growing period in the growth chamber was considered to be relatively short for the seedlings to become nutrient deficient. At planting time seedlings were watered to field capacity (900 mL of water/pot). Thereafter, seedlings were watered every 3-4 days (50 mL/pot) or whenever necessary to maintain the root medium at field capacity. Growth chamber conditions during the post-storage period were as follows: temperature 11 ° C day and night, relative humidity 70%, photoperiod 16 hours, photosynthetic photon flux density 350 gmol quanta m -2 s -1 , from cool white fluorescent lights (Philips VHO), supplemented with incandescent bulbs. Seedlings were grown for 28 days during which time bud break assessments were conducted every two days. On the 28th day, seedlings were removed from the growth chamber, their root medium carefully washed off, and  21  new white roots counted (i.e. new roots > 5 mm long).  Figure 2. A general view of the pot arrangement of the CE  experiment in the growth chamber in one of the water baths.  22  3.1.3. Carbohydrate Analyses  At each planting time, some seedlings were randomly selected (all six CO 2 treatments represented) and harvested for analysis of starch and soluble sugar content. The harvested seedlings were divided at the root collar into roots and shoots, killed by freezing them in liquid nitrogen and then they were freeze-dried for 3-4 days. Together with the previously harvested samples, starch, soluble sugars, and their sum, total non-structural carbohydrates (TNC), were determined by methods modified from da Silveira et a/. (1978), Haissig and Dickson (1979), Jermyn (1975), Rose et al. (1991), and Yemm and Willis (1954). Seedlings from the first set of reciprocal transfer experiments (60 days after sowing) were included in the analysis only at 80 and 120 days. For the entire experiment, a total of 1284 carbohydrate samples were analyzed.  3.1.3.1. Soluble Sugars Analysis by Anthrone Reagent  Soluble sugars (i.e. free sugars), expressed as glucose equivalents, were determined by methods modified from Jermyn (1975), and Yemm and Willis (1954). Details of the analytical procedure are reported in Appendix 12 (item V). The already freeze-dried plant material (i.e. needles, stems, and roots) was finely ground in liquid nitrogen with a mortar and pestle. Twenty-mg samples were extracted with five mL methanol:chloroform:water (M:C:W) (12:5:3, v/v/v) overnight. After centrifugation (desk top) the pellet was  23  re-extracted with three mL M:C:W. Supernatants from the two extractions were then combined, and three mL of distilled water were added. After centrifugation, the aqueous phase was removed and evaporated to dryness. Thereafter, the dried sample was dissolved in 3 mL of distilled water and stored frozen until required for sugar analysis. Prior to analysis, 200 gL samples were treated with 50 gL each of Ba(OH)2 and ZnSO4 (0.3N) to remove organic acids and proteins, and soluble sugars were analyzed by anthrone according to Jermyn (1975), and Yemm and Willis (1954).  3.1.3.2. Starch Analysis by Enzymatic Hydrolysis  Starch was analyzed by gelatinizing the pellet from the two extractions above in five mL of acetate buffer (150mM, pH 4.5) and autoclaving at 120 ° C for one hour. After the mixture was cooled to 55 ° C in a water bath, 125 units of amyloglucosidase (E.C. 3.2.1.3; from Rhizopus, Sigma A-7255) and 25 units of a—amylase E.C. 3.2.1.1 (from Aspergillus, Sigma A-0273) were added to the samples, and the mixtures were then incubated for two hours. Thereafter, the mixtures were centrifuged (desk top), and the supernatants analyzed for starch (expressed as glucose equivalents) using glucose oxidase-peroxidase-o-dianisidine (Ebell 1969, Haissig and Dickson 1979, Rose et a/. 1991). For more details on the analytical procedures, refer to Appendix 12 (item IV).  24  3.2. Experiment Two: Stable Carbon Isotopes as Tracers 3.2.1. Seedling Production and Cold-storage  The second experiment involved producing Engelmann spruce seedlings (the same seediot as in the first experiment) in 213 styrofoam blocks at the UBC South Campus Nursery beginning in May, 1990. The standard nursery procedures were followed up to bud set (September). Thereafter, seedlings were moved into a Conviron E30 growth chamber, and placed in four water-cooled 125 dm 3 Plexiglas boxes for stable carbon isotope "labelling" (Fig. 3). Each box contained 50 seedlings; two boxes acted as controls (i.e. receiving ambient CO2 with a normal isotopic composition) and the other two were "labelled" (i.e. receiving air that was first stripped of ambient CO2, before adding back CO2 depleted in  13 C).  A computer data  acquisition and control system (WB 820 Omega Engineering, -  Inc., Stamford, CT) and mass flow controllers (MFC 825, Edwards High Vacuum Intl, Wilmington, MA) matched CO2 concentrations in "labelled" boxes with the "unlabelled" boxes by injecting tank CO2 into the former. The IRGA (LI6251 CO2 Analyzer from LI-COR, Lincoln, NB) was used to monitor CO2 concentrations in the boxes. The Plexiglas boxes were provided with small fans and de-humidifying air recirculation loops. Tank CO2 going into "labelled" boxes had a 5 13 C value of -35.81%. Initial growth chamber conditions were as follows: day and night temperatures 22/15 ° C, photoperiod 12 hours,  25  Figure 3. A schematic diagram of equipment used to produce seedlings differing in stable carbon isotope composition (isotope labelling experiment). Two of four boxes are shown. Air taken from outside was split into two streams, and one stream was pumped directly into control (i.e. "unlabelled") boxes. The second stream was stripped of CO2 by passing through soda lime before being pumped into boxes for the "labelled" seedlings. An infra-red gas analyzer (IRGA) was used to monitor CO2 concentrations. A computer data acquisition and control system and mass flow controllers (MFC) matched CO2 concentrations in the "labelled" boxes with the "unlabelled" boxes by injecting tank CO2 into the former. The computer also monitored and controlled the air temperature and humidity in the boxes.  26  ^ Air OUT  Pump "unlabelled"  IRGA  Outside Air IN = —8 %.  ■••■•■•■■  Soda Lime  .■•■11.  Filter  Pump  •■■•=01•4  !44 't I 1  "labelled"  MFC Tank CO 2 (5 13C = —36 7«,  Air OUT  27  relative humidity 50% during the day and 60% at night, and photosynthetic photon flux density ca. 300 lamol quanta m -2 s -1 . On November 30, 1990, the temperature in the growth chamber was reduced to 10/5 ° C day/night in order to hardenoff the seedlings before putting them into cold-storage. Seedlings were watered to field capacity every three days (i.e. 2.5 L of water per box, allowed to freely drain for 10 minutes). Furthermore, 20:8:20 N:P:K fertilizer was applied to the seedlings once a week for the first four weeks and then once every fortnight thereafter. On December 19, 1990, seedlings were lifted, packed in plastic-wrap, placed in lined boxes and taken into coldstorage. At the same time, six seedlings were harvested from each treatment (i.e. control and labelled). The harvested seedlings were then killed in liquid nitrogen, freeze-dried, and stored at -5 ° C until required for carbohydrate and stable carbon isotope analyses.  3.2.2. Post Cold-storage Planting  For the carbon isotope tracer study, seedlings were withdrawn from the cold-storage for planting in the growth chamber at the UBC South Campus Nursery only once, in April, 1991. Growth chamber conditions and root medium were as described in section 3.1.2. However, here 30 pots were placed in each water bath (three water baths all at 11 ° C) and only one seedling was placed in each pot. Hence, there were 15 pots for the "labelled" seedlings and 15 pots for  28  control seedlings, all randomly placed in each water bath (Fig. 4).  (A)  (B)  Figure 4. General views of the equipment arrangement when  seedlings were moved into the Plexiglas boxes for (a) isotope labelling, and (b) during post-storage planting.  29  Also at planting, six seedlings from each treatment were harvested and preserved for carbohydrate and isotope analyses. Seedlings were grown in the growth chamber for 36 days. On May 9, 18, and June 5, 1991, five seedlings for each treatment were harvested from each water bath. The root medium was carefully washed off the harvested seedlings, new white roots collected, killed in liquid nitrogen, freezedried, and then stored in the freezer until required for stable carbon isotope analysis.  3.2.3. Stable Carbon Isotope Analyses  Before the new roots were analyzed for 13 C/ 12 C ratio, the already harvested pre- and post-cold-storage samples were analyzed for carbohydrate reserves. The analytical procedures in chapters 3.1.3.1 and 3.1.3.2 were followed. The carbohydrate analysis at this stage was necessary only for establishing the level of carbohydrate reserves in the plant material we were dealing with. In addition, before the actual 13 C/ 12 C ratio analysis, samples (new white roots, starch and soluble sugars extracts) were prepared. The samples were finally assessed for 13 C/ 12 C ratio using the methods adapted from Boutton (1991), Deleens et al. (1989), Engel and Maynard (1989), Guy and Wample (1984), and Svejcar et al. (1990). In essence the methods involve changing the plant material into CO2 by combustion for determination of the 13 C/ 12 C ratio by mass spectrometry. The isotope analysis  30 was performed by the Isotope Mass Spectrometry Laboratory, Department of Oceanography, using a VG Isogas Prism triplecollecting mass spectrometer. The 8 13 C was calculated from the measured carbon isotope ratios of the sample and standard gases as:  813 CW  = [(Rsample-Rstandard)/Rstandard] x 10 3  where 8 13 C is the parts per thousand, or per mil (tD), difference between 13 C content of the sample and the standard, and R is the mass 45/44 ratio of sample or standard gas. The 8 13 C values were expressed relative to the international PDB (Pee Dee Belemnite) standard. Sample preparation and machine precision was ± 0.1% -o. The fractional abundance (F) was estimated by the following equation:  F^(RL-RC)/(CL-CC)  where F is fraction of old carbon, R L is current labelled root 8 13 C, RC is current control root 8 13 C value, CL is labelled old carbon source (i.e. carbohydrate) 5 13 C, and CC is control old carbon source (i.e. carbohydrate) 8 13 C value. The values used for C L and CC are weighted means of separate 8 13 C values for sugars and starch.  31 3.2.3.1. New White Roots  Due to the small amounts of tissue involved, the freeze-dried root samples were not ground or pulverized in a conventional way (i.e. grinding with a mortar and pestle or in a Wiley mill). Instead, these were ground as finely as possible within their storage vials using a clean glass rod. The ground samples were kept in a desiccator under vacuum until required for combustion. Combustion tubes were prepared from quartz tubing (6 mm o.d. x 4 mm i.d.) cut into 25 cm lengths, fused shut at one end. Tubes were loaded with ca. 1.55 g of cupric oxide wire (pre-fired at 550 ° C), 2.0 g reduced copper and plant sample (3 - 10 mg). The sample tubes were then attached to a vacuum line, evacuated to less than 0.13 Pa (10 -3 torr), and sealed with a torch. Sealed sample tubes were placed in a muffle furnace and combusted at 900 ° C for two hours to convert all the organic carbon to CO2 quantitatively. The combusted sample tubes were stored at room temperature for a month, but were reheated to 550 ° C for ca. one hour just prior to isotope analysis. Reheating of samples was necessary because Engel and Maynard (1989) found that, if combusted tubes are stored for more than five days before analysis, CO2 becomes depleted by 1-3% as a result of carbonate formation. If samples cannot be processed within five days, this isotopic depletion can be overcome by recombusting the tubes prior to the actual isotope analysis (Engel and Maynard, 1989).  32  3.2.3.2. Starch Extraction  Six 100 mg samples of the already ground plant material from each treatment were pooled together, representing four 600 mg samples (i.e. two for pre- and post-cold-storage treatments respectively). Separation of soluble sugars from starch and the preliminary extraction of soluble sugars were done following the procedures outlined in Appendix 12 (items II and III). However, in this case, volumes for M:C:W and distilled water were increased four-fold to account for the increased amount of plant material. The starch pellet recovered was then freeze-dried, and starch was finally extracted by methods adapted from Brugnoli et al. (1988), Ehleringer (1991), and Hassid and Abraham (1957). Two hundred and fifty mg of the starch pellet was weighed out and placed in a 25-mL centrifuge tube. Then 25 mL of boiling 80% ethanol were added and the mixture boiled in a water bath for 15 minutes. After cooling, the samples were centrifuged (desk top) and the supernatant discarded. The extraction was repeated until the supernatant was colourless. The pellet was re-suspended in 20 mL of 20% hydrochloric acid (w/w) for 30 minutes, spun down (desk top) and the supernatant saved. The pellet was re-extracted with 20 mL of 20% hydrochloric acid and spun down as before. The two supernatants were then combined in a volumetric flask, made up to 100 mL with distilled water, and then 6.6 g of sodium chloride and 10 mL of 0.14N iodine-potassium iodide solution were added and left standing on ice for 45 minutes.  33  Thereafter, the precipitate was centrifuged in 250 mL centrifuge bottles at 1000 g for 10 minutes at 5°C (Beckman JA-14 rotor). The supernatant was discarded. The pellet was then re-suspended in alcoholic sodium chloride (0.34N in 70% ethanol) (4x5 mL) for transfer to a 25 mL centrifuge tube, spun down (desk top), and the supernatant discarded. Four mL of 0.25N alcoholic sodium hydroxide was added to the pellet and left in the refrigerator overnight (repeated where necessary until the brownish colour disappeared). The following day, the alcoholic sodium hydroxide was poured off, the starch pellet was re-suspended (2x) in 15 mL of 60% ethanol, centrifuged (desk top), and the pellet saved. The starch pellet was then dissolved in 10 mL of distilled water by autoclaving at 120 ° C for one hour. The aliquot of starch extract was filtered, and then reprecipitated by adding 15 mL of 100% ethanol. The precipitation did not occur spontaneously, and so another 10 mL of 100% ethanol were added and the samples were then kept in the freezer for approximately two days. Thereafter, the precipitate was centrifuged at 12,000 g (Beckman JA-20 rotor) for 20 minutes at 5 ° C, and the supernatant discarded. Fifteen mL of 60% ethanol were added to the pellet and allowed to sit for five minutes. The fluid was discarded and rinsing was repeated with 95% ethanol, 100% ethanol, and then ether (2x). The starch extract was finally air-dried at room temperature for ca. 30 minutes, and the samples stored in a desiccator until needed for isotope analysis.  34  3.2.3.3. Soluble Sugars Extraction  Methods described by Brugnoli et al. (1988) and Ehleringer (1991) were adapted to extract soluble sugars. First of all, resins (Dowex-50 (11 + form) and Dowex-1 (C1 -1 form)) were prepared as follows: Both resins were washed with distilled water several times to remove fines and bring the slurry pH to that of distilled water (ca. 6.0). For activation, 10-fold excess of 1N HC1 and 2N HC1 were added to Dowex-50 and Dowex-1 respectively. These were allowed to stand, with occasional stirring, for 15 minutes. Thereafter, the slurry was filtered, and washed well with distilled water until the pH was approximately 6.0. Five mL of each resin were loaded into separate 10-cc Luer-Lock syringes. These "columns" were arranged in series with the Dowex-1 on top, and the Dowex-50 below. Aqueous fractions obtained from the preliminary soluble sugars extractions (12 mL per treatment) were then loaded onto the columns and eluted with 180 mL of distilled water. The eluate was then filtered, and flash evaporated to a final volume of two mL. The samples were re-filtered, freezedried, and then stored in a desiccator under vacuum, until required for isotope analysis.  35  3.3. Experimental Design and Data Analysis 3.3.1. Experiment One  The experimental design was divided into two parts. The first part investigated carbohydrate reserves and seedling growth prior to cold-storage, in which case, a completely randomized design was used (Appendices 1-10) Six dates (i.e. at 80, 120, 140, 202, 268, and 327 days) were considered, and at each date only the CO2 treatment effects were statistically analyzed. Time, as a factor, was not included in the analysis. There is ample evidence in the literature indicating that carbohydrate levels in conifers fluctuate seasonally and overall trends observed in the present study were as expected. Four treatments were considered: low, high, step-down, and step-up CO2 concentrations. The number of replicates for each treatment varied considerably from one month to another; with n=12 for 80, 120, and 140 days, n=7 for 202 days, n=2 or 6 (depending on treatment) for 268 days, and n=6 for 327 days. The second part of Experiment One involved assessment of bud break and root growth at 268 and 327 days. The experimental design at these two dates was similar, except for a slight modification at 327 days. At 268 days a completely randomized block design with a 2 x 2 factorial arrangement (i.e. six CO2 treatments and three water bath temperatures) was used (Appendix 11). However, it is worth mentioning here that, since only one water bath was used for each water bath temperature for the entire growth period,  36  there was no real replication of the water bath temperatures. According to Hurlbert (1984), this situation can be viewed as pseudo-replication. In the context of this study, pseudo-replication may refer to the testing for water bath temperature effects without true replication of the treatments. In this study the assumption was made that the growth chamber environment was reasonably uniform and that water baths differed only with respect to temperature. The number of replicates for each treatment remained fairly constant (n=6) from one water bath to another, except for a few situations with n=5 or 7. At 327 days, the same design used for 268 days was applied. However, in this case, all the three water baths were at 11 ° C. The number of replicates (n=6) for each treatment remained consistent from one water bath to another. The statistical analysis involved analysis of variance, and the computer statistics program, Systat, version 5.0 (Wilkinson 1990) was used to do the analysis. The significance was tested at the P<0.01 level. The 1% level of significance was used in order to account for multiple and post hoc hypotheses, based on the Bonferroni inequality procedure (Meddis 1984, Wilkinson 1990). Group variances were tested for homogeneity using Bartlett's test (Walpole 1982). Every ANOVA had homogeneous variances. Where the analysis of variance showed significant differences, mean separation was accomplished by Tukey's test (Ott 1984, and  37  Zar 1984) at the P<0.01 level.  3.3.2. Experiment Two  As in 3.3.1, the experiment was divided into two parts. Part one of the experiment covered pre- and post-cold storage data for carbohydrate reserves. A completely randomized design with a 2 x 2 factorial arrangement (i.e. two storage periods and two treatments: control and "labelled") was used. The control and "labelled" treatments were replicated six times (n=6). The second part of Experiment Two covered data for the stable carbon isotope analysis. The initial intention was to have a completely randomized block design, representing two treatments and three water baths (all water baths at 11 ° C). However, due to low rooting capacity by the seedlings it was not possible to perform a two way analysis of variance. Instead the data were subjected to a t-test, with n=9 for the second harvest, and n=6 for the third harvest. For the first harvest only one "labelled" seedling produced some new white roots, hence it was not possible to perform any statistical analysis on a single observation. The computer package used in 3.3.1 for the statistical analysis was applied here too. The significance was tested at the P<0.01 level.  38  4.0 RESULTS 4.1 Carbon Dioxide Enrichment Experiment 4.1.1 Effects of CE on Growth Prior to Cold-storage 4.1.1.1 Biomass Accretion and Partitioning  Biomass accretion in individual tissues within seedlings, in response to CE, varied greatly at different stages of seedling development (Figs. 5 and 6a). From Figs. 5 and 6a it can be noted that, CE significantly (P<0.001) affected needle and stem biomass at 80, 140, and 202 days, and root biomass at 80 and 140 days. In all the individual plant parts, there was a general positive response in biomass production to high CO2 and step-up treatments relative to low CO 2 concentration. The step-down treatment had a negative response relative to its control (i.e. the high CO2 treatment) but still showed greater final biomass production than the continuous low CO2 treatment. Even where there were no significant differences between treatment means, high, step-down, and step-up CO2 treatments generally recorded higher dry weights than the low CO2 treatment. At the end of the growing season (202 days), needles  recorded the highest biomass followed by roots, and the stems recorded the least. In all the individual plant tissues, the most significant effects of CE were after bud  set (140 and 202 days). Because after bud set, shoot height growth had almost ceased, increased biomass at this stage of seedling production could be attributable to increased  39  girth, root growth or reserve accumulation (mostly sugars and starch). Total seedling biomass followed about the same trend as the individual tissues (Fig. 6b). At all stages of seedling development, except at 120 days, CE significantly (P<0.001) affected total seedling biomass. Here also, the pattern of high CO 2 levels being superior to low CO2 levels was prevalent. It is also important to note that, after bud set, both individual tissues and total seedling biomass for enriched seedlings were almost two-fold greater than for seedlings grown at ambient CO2 levels. The shoot:root ratios (on a dry weight basis), decreased steadily during the whole study period and were not generally affected by CE (Fig. 7). This pattern of decreasing shoot:root ratios, especially after bud set was as expected because at this stage of development the shoot sink strength would be minimal owing to cessation of shoot height growth. The significant (P<0.001) effect of CE observed for the step-down treatment at 120 days stands out but does not conform to any general pattern, and is without obvious explanation.  40  Figure 5. Effects of CE on (a) needle dry weight, and (b) stem  dry weight of Engelmann spruce seedlings. The value for each bar is a mean of 24 seedlings. Step-down and step-up treatments represent seedlings that were transfered between CO2 environments at 60 and 120 days. Step-up and step-down data presented for 80 and 120 days are for transfers performed at 60 days, while further data are for transfers performed at 120 days. At each age, bars accompanied by the same letter(s) or no letters were not significantly different at the P<0.01 level.  41  (A)  1200  1000  E  111/ Low CO 2 11 High CO 2 N'S1 Step—down ES) Step—up iSE  b  b  800  rn 600  ID^400  w  co 200  80^120^140^202  Age (days)  (B)  600 11111 Low CO 2 7 1 High CO 2 ^ Step—down 22 Step—up -  500  b  ±SE 400  300  200  100  80^120^140  Age (days)  ^  202  42  (A) 1000 ^  800  MI Low CO 2 High CO 2 1['Q Step-down (a3 Step-up *SE  600 b  400  200 ab  a  •  14  (B) 2500 ^  2000  Cn E a)  •  0  NMI Low CO 2 ri High CO 2 17KI Step-down Egl Step-up  b  *SE b  1500  1000  0 .  0 -  b  500  ab  80^120^140  ^  202  Age (days) Figure 6. Effects of CE on (a) root dry weight, and (b) total  seedling biomass of Engelmann spruce seedlings. The value for each bar is a mean of 24 seedlings. At each age, bars accompanied by the same letter(s) or no letters were not significantly different at the P<0.01 level. For details see Fig. 5.  43  5  EN  Low CO 2 High CO 2 NN1 Step-down 22:1 Step-up -  4  ±SE  CI 0  3  2  1  80^120^140  ^  202  Age (days) Figure 7. Effects of CE on shoot:root ratio (DW basis) of Engelmann spruce seedlings. The value for each bar is a mean of 24 seedlings. At each age, bars accompanied by the same letter(s) or no letters were not significantly different at the P<0.01 level. For details see Fig. 5.  44  4.1.1.2 Root Collar Diameter and Stem Height  Root collar diameter of Engelmann spruce seedlings responded positively to CE at all stages of seedling development except at 120 days (Fig. 8a). As in the case of biomass, the response to CE was more significant (P<0.001) after bud set (140 and 202 days), with high, step-down, and step-up CO2 treatments being generally superior to low CO2 treatment. Note the steady increase in root collar diameter after bud set, particularly in seedlings grown under high CO2 regime. This steady increase in root collar diameter after bud set and the cessation of shoot height, indicates that a portion of the available assimilates from continuing photosynthesis were used in radial growth. In other words, in terms of size, the main effect of CE after bud set is that seedlings became "stockier". The effects of CE on stem height of Engelmann spruce seedlings are presented in Fig. 8b. Data reported here are only for seedlings grown under high and low CO2 concentrations (i.e. controls) Data for step-down and stepup treatments were omitted because the trends were similar to controls. Data for 202 days were not taken because shoot height growth had ceased long before this date. Carbon dioxide enrichment significantly (P<0.001) affected stem height only at 140 days. However, at all previous harvests, even before 80 days (data not presented), high CO 2 seedlings always had a greater mean height than low CO 2 seedlings.  45  (A) 3.0  ,—. E 2.5 E L.  w 2.0 i  .4  E 0 6 1.5 L.  0  o 1.0 C.)  "6 0 ix 0.5  0.0  80^120^140  ^  202  Age (days) (B) 14 12  11111 Low CO 2 I^I High CO 2 tSE  I  b  80^120^140  Age (Days)  Figure 8. Effects of CE on (a) root collar diameter, and (b) stem  height of Engelmann spruce seedlings. The value for each bar is a mean of 24 seedlings. At each age, bars accompanied by the same letter(s) or no letters were not significantly different at the P<0.01 level. For details see Fig. 5.  46  Overall, it appears that CE had a greater effect on biomass production than on stem height and root collar diameter of Engelmann spruce seedlings. 4.1.2 Carbohydrate Reserves  There was a general tendency for carbohydrate levels to decrease initially at 120 days, presumably due to the reduction in photoperiod (Figs. 9-16). Following this, as shoot growth was curtailed, reserves began to accumulate. Carbon dioxide enrichment significantly (P<0.001) increased the whole plant total non-structural carbohydrates (TNC) only after bud set (120 and 140 days in Fig. 9). Both the high and step-up CO2 treatments showed this increase in TNC. However, by the time seedlings were ready for cold-storage, TNC levels were the same in all treatments. After two and four months (at 268 and 327 days, respectively) of storage, there was almost a one-third reduction in reserves. The reduction in TNC during the two storage durations was comparable, although the depletion was slightly more after four months storage. In terms of reserve allocation, roots contained about 50% of the whole plant TNC just before storage (202 days), whereas, at all other times, needles had the highest  47  Whole Plant 500  400  ami Low CO  Cold—storage  9—h day  18—h day I  2  High CO 2 Step-down 1\\\1 EEO Step-up ±SE  b b  c) 0) 300 0)  E  200 F1 00  80^120^140^202  268  327  Age (days) Figure 9. Trends in whole plant total non-structural  carbohydrates (TNC) of Engelmann spruce seedlings. The value for each bar is a mean of 2-6 (at 268 days), 6 (at 202 and 327 days), and 12 (at 80, 120, and 140 days) seedlings. At each age, bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  48  Reserve Allocation 18—h day I  eTh  0  9—h day  Cold—storage  75  E  4 0  -  RI 50  F—  80^120^140^202  268  327  Age (days) Figure 10. Trends in allocation of reserves to individual plant  parts, expressed as a percentage of the maximum total nonstructural carbohydrates (TNC) observed just prior to storage (i.e. at 202 days). At each age, the proportion is a total of all treatments representing 16 (at 268 days), 24 (at 202 and 327 days), and 48 (at 80, 120, and 140 days) seedlings.  49  proportion (Fig. 10). However, by the end of the two storage durations, there was a considerable reduction (more than 50%) in root TNC, and the needles at these times, constituted more TNC than any other plant part. Stems recorded the lowest allocation of reserves during the whole study period. Like needles, stem TNC percentages remained fairly consistent. Respiration was presumably not restricted to the roots and, in addition to the depletion of TNC during storage as an energy source for root respiration and maintenance, it is logical to suspect that some root TNC might have been translocated out of the roots (after starch conversion to glucose) to the needles and stems. During the whole study period, CO2 treatments significantly (P<0.001) affected soluble sugar content in needles at 140 days (Fig. 11), in stems at 80 days (Fig. 12), and in roots at 80, 120 and 202 days (Fig. 13). Throughout the whole study period, particularly after bud set, there was a general trend for high and step-up CO2 treatments to contain more soluble sugars than the others. Overall, needles had the highest accumulation of soluble sugars, while stems and roots accumulated lower but comparable amounts. In this study, it appears that all the three plant parts of Engelmann spruce seedlings accumulated and stored appreciable amounts of soluble sugars. For the needles, soluble sugar content ranged from approximately 200 to 350 mg g -1 DW, with 140 and 202 days recording the  50  Needles 18—h day I  500  um  ^  9—h day  ^  Cold—storage  Low CO 2 High CO 2  RN 0 400  Step-down  EKg Step-up ±SE  b  b  cn 300 rn  cn  7  ur) w  200  D  6 100  -  yl  80^120^140^202  ^  268  ^  327  Age (days) Figure 11. Trends in soluble sugar content of needles (buds included) of Engelmann spruce seedlings. The value for each bar is a mean of 2-6 (at 268 days), 6 (at 202 and 327 days), and 12^(at^80,  120, and 140 days) seedlings. At each age,  bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  51  Stems 18—h day I  500  9—h day  Cold—storage.  Low CO 2 ^ High CO 2 ^ Step-down  0 400  EZ Step-up ±SE  cn cn 300  cn = ul 200 0 .15 6 100  -  0  80^120^140^202  268^327  Age (days) Figure 12. Trends in the soluble sugar content of stems of Engelmann spruce seedlings. The value for each bar is a mean of 2-6 (at 268 days), 6 (at 202 and 327 days), and 12 (at 80 days, 120 days, 140 days) seedlings. At each age, bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  52  Roots Cold  18—h day^9—h day  500  —  storage  Low CO 2  High CO 2  N\1 C) 400  EDE]  Step-down Step-up  ±SE  Cr)  b  300 0  rn Ul  200  0  75 1 00  80  ^  120^140^202^268^327  Age (days) Figure 13. Trends in soluble sugar content of roots of Engelmann  spruce seedlings. The value for each bar is a mean of 2 (at 268 days), 2-6 (at 202 and 327 days), and 12 (at 80, 120, and 140 days) seedlings. At each age, bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  53  highest values (Fig. 11). Interestingly, there was no major depletion of soluble sugars in the needles after two and four months cold-storage durations. In the case of stems and roots, soluble sugar content ranged from about 90 to 250 mg g -1 DW and the proportions in these plant parts were fairly close to each other at most times (Figs. 12 and 13). Soluble sugar content in the stems was not affected by the two cold-storage durations, whereas in the roots there was a reduction of about 20%, and this reduction was almost the same at both durations. In the case of starch, CO2 treatments had a significant (P<0.001) effect on needles, stems, and roots only at 140 days (Figs. 14-16). Here also, both the high and step-up CO2 treatments showed this increase in starch. Contrary to the trends observed in soluble sugars, starch accumulation and storage did not seem to take place equally in all the three plant parts studied. Instead, roots were the major storage organ for starch, followed by stems (Figs. 15 and 16). In both roots and stems, starch accumulation rose sharply after bud set, the period during which CE was probably more effective. From Figs. 12 and 13, it can be noted that, during the earlier stages (e.g. 80 and 120 days), starch accumulation in stems and roots of Engelmann spruce seedlings was minimal. As for soluble sugars, maximum starch accumulation in  54  Needles 18—h do  500  in  9—h day  Cold—storage  Low CO 2 High CO 2  N\1 400  Step-down  1221 Step-up ±SE  c) cn  300  Cr)  E  0 200 65 -  100  80^120^140^202  ^  268  ^  327  Age (days)  Figure 14. Trends in starch content of needles (buds included) of  Engelmann spruce seedlings. The value for each bar is a mean of 2-6 (at 268 days) , 6 (at 202 and 327 days) , and 12 (80, 120, and 140 days) seedlings. At each age, bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  55  Stems 18—h day I  500  IN 1  ^  9—h day^i^Cold—storage  Low CO 2 High CO 2  r\\] Step-down  400  [Kg Step-up ±SE  c) cn  ci)  300  E  0 200  100  80^120^140^202  ^  268  ^  327  Age (days)  Figure 15. Trends in starch content of stems of Engelmann spruce  seedlings. The value for each bar is a mean of 2-6 (at 268 days), 6 (at 202 and 327 days), and 12 (80, 120, and 140 days) seedlings. At each age, bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  56  Roofs 500  as Low CO I^ I  \I  400  Cold—storage  9—h day  18—h daY 2  High CO 2 Step-down  EZ Step-up ±SE  cy)  30 0  cn  E  _c  0  b  200  b  100  M  IMI IMO  80^120^140^202  ^  268  ^  327  Age (days)  Figure 16. Trends in starch content of roots of Engelmann spruce seedlings. The value for each bar is a mean of 2-6 (at 268 days), 6 (at 202 and 327 days), and 12 (at 80, 120, and 140 days) seedlings. At each age, bars accompanied by the same letter(s), or no letters were not significantly different at the P<0.01 level.  57 roots (ca. 500 mg g -1 DW) and stems (ca. 200 mg g -1 DW) was observed just before storage (202 days). In needles, the maximum accumulation of starch was observed at 80 days, and was less than 100 mg g -1 DW (Fig. 14). The results vividly show that, for Engelmann spruce seedlings, needles do not function as a major storage organ for starch. In all the individual plant organs, there was more than 50% reduction in starch content after two and four months storage, with the needles recording only a minimal amount. 4.1.3 Bud break and Root Growth Potential  In this study, bud break was assessed as mean days to first bud break. None of the CO2 treatments had any significant effect on days to first bud break at either 268 or 327 days at the P<0.01 level. At both ages, there were no logical trends observed for the treatment effects. However, at 268 days, where soil temperature was considered as a factor, bud break was significantly (P<0.001) affected, with the 3 ° C soil temperature treatment recording a greater number of days to first bud break (ca. 15 days) than the 7 and 11 ° C soil temperature treatments (Table 1). On average, the lowest number of days to first bud break (ca. 10-12 days) was observed at 327 days (Table 1). At 327 days, days to first bud break were generally fewer than at 268 days even when seedlings were grown at the same soil temperature (e.g. 11 ° C). The fewer days to first bud break observed at 327 days could be explained by the fact that, with longer  58  Table 1. The effects of pre-storage CE on days to first bud  break of Engelmann spruce seedlings. Values are treatment means of 6 seedlings (at 268 days) and 18 seedlings (at 327 days). Values in brackets are standard errors. Step downl/step upl and step down2/step up2 represent seedlings transfered at 60 and 120 days, respectively. Treatment means at each soil temperature were not significantly different at the P<0.01 level.  At 268 days Soil Temp.  Low  3°C  High  Step downl  14.8 (0.49)  13.33 (0.67)  14.29 (0.29)  14.33 (0.33)  15.14 (0.59)  15.0 (0.68)  7° C  13.67 (0.33)  13.71 (0.72)  12.67 (0.42)  12.56 (0.44)  14.25 (0.25)  14.33 (0.33)  11 ° C  13.77 (0.36)  13.5 (0.72)  13.33 (0.42)  13.0 (0.45)  13.67 (0.33)  13.35 (0.44)  Step upl  Step down2  Step up2  At 327 days 11 ° C  9.67 (0.74)  11.57 (0.95)  11.89 (0.42)  10.89 (0.57)  12.56 (0.79)  11.35 (0.57)  59  Table 2. The effects of pre-storage CE on post-storage RGP (number of new roots >5 mm long) of Engelmann spruce seedlings. Values are treatment means of 6 seedlings (at 268 days) and 18 seedlings (at 327 days). Values in brackets are standard errors. Step downl/step upl and step down2/step up2 represent seedlings transfered at 60 and 120 days, respectively. Treatment means at each soil temperature were not significantly different at the P<0.01 level.  At 268 days Soil Temp.  Low  High  Step downl  Step upl  Step down2  Step up2  7 °C  14.17 (6.17)  15.50 (5.82)  20.17 (12.39)  18.50 (9.18)  22.67 (11.64)  24.16 (11.64)  11 ° C  20,16 (6.99)  64.5 (22.36)  53.00 (24.77)  47.83 (10.37)  51.33 (9.28)  68.67 (10.89)  66.11 (8.38)  58.26 (10.08)  60.39 (10.00)  At 327 days 11 ° C  42.11 (8.39)  27.78 (5.97)  47.22 (11.48)  60  cold-storage durations, seedlings had a greater opportunity to attain the necessary chilling, and thereafter, accumulate adequate heat units to permit the resumption of shoot growth. Similarly, at both 268 and 327 days, RGP was not significantly affected by any of the CO2 treatments at the P<0.01 level (Table 2). However, at both ages, the low and high CO2 treatments consistently showed low RGP values relative to step-down and step-up CO2 treatments. As for bud break, at 268 days, where soil temperature was considered as a factor, root growth was significantly (P<0.001) affected, with the 7 ° C soil temperature treatment recording extremely low numbers of new roots (ca. 10-20) as compared to the 11 ° C soil temperature treatment (ca. 50-70) (Table 2). Unlike bud break, RGP values for 268 and 327 days at 11 ° C soil temperature were comparable. 4.2 Stable Carbon Isotopes as Tracers  Whole seedling isotope composition and carbohydrate reserves before and after cold-storage are reported in Table 3. The TNC in unlabelled and labelled seedlings were nearly the same, implying that the procedures used for isotope labelling did not differentially affect the carbohydrate status of the seedlings. As with the CE experiment, carbohydrates were depleted by approximately 50% after four months cold-storage.  61  Table 3. Isotopic composition (expressed as 8 13 C value) of whole  seedling total tissue and total nonstructural carbohydrates (TNC) before and after cold-storage.  8 13 C values (^) Starch^Sugars  Total tissue  TNC (mg/g DW) ±SE  Pre-cold storage Unlabelled  -21.89  -23.77  -25.25  394 ±39  Labelled  -34.09  -40.38  -33.43  439 ±23  Unlabelled  -21.44  -24.99  -25.57  199 ±27  Labelled  -35.22  -38.84  -33.01  237 ±22  Post-cold storage  62  Table 4. Trends in isotopic composition (expressed as 8 13 C  value) of new white roots, and calculated per cent contribution of reserve carbon to new root construction of "labelled" seedlings.  Days after planting^Mean 8 13 C (to) ±SE^% reserve carbon Unlabelled^Labelled^±SE (N) 9 d  *  -36.78  100 (1)  18 d  -22.94 ±0.64  -30.26 ±1.02  52.9 ±6.9 (9)  36 d  -21.91 ±0.41  -24.47 ±0.76  18.3 ±5.0 (6)  * Control seedlings at this date did not have any new roots.  63  As desired, there were clear differences between unlabelled and labelled seedlings in starch, sugar, and total tissue 513 C values. There were also differences in  813 C of total tissue, but these were not as marked because unlike the bulk of the TNC, total tissue would contain large amounts of carbon fixed prior to the labelling period. Futhermore, for each treatment, the 513 C values were virtually the same before and after cold-storage, indicating that cold-storage did not alter isotopic composition of the seedlings. Table 4 shows trends in the isotopic composition of new white roots, and calculated contributions of reserve carbon to new root construction. Results from this study indicate that after 9 days, 18 days, and 36 days, planted Engelmann spruce seedlings contained respectively, 100, 53, and 18% old carbon in the new roots. However, these contributions represent only a very small proportion of the available reserves remaining after storage. This is because biomass of new roots (ca. 3.8 mg/seedling) was small relative to the size of the reserve carbohydrate pool (ca. 237 mg/seedling after storage). Assuming new roots are composed principally of carbohydrates (including cellulose), and are therefore about 40% carbon by dry weight, and knowing that after 36 days, 18% of this carbon originated from reserves, it can be calculated that less than 1% of the reserve pool ended up in these roots (only ca. 0.3% in fact). It is however, not  64  known where or how the rest of the reserve carbon was used. It is possible that some of the old carbon remained unused. Otherwise, much of it could have been, (1) used in root respiration during the construction of new roots, (2) translocated to other growing tissues (e.g. cambium, shoot meristems) and used in the resumption of shoot growth, or (3) used in various metabolic processes for repair and/or maintenance of extant tissues.  65 5.0 DISCUSSION 5.1 Biomass Accretion and Partitioning  This study has revealed that, elevated CO2 levels influenced biomass accretion in individual tissues of Engelmann spruce seedlings, and that these effects were more distinct after bud set (Figs. 5 and 6a). In all treatments, the greater allocation of biomass to needles and/or leaves followed by roots in Engelmann spruce seedlings in this study is in agreement with seedling biomass accretion for other species (Brown and Higginbotham 1986, Campagna and Margolis 1989, Hagem 1948, Sionit et al. 1985, and Hollinger 1987). Similarly, the trends in total seedling biomass were identical to those for the individual plant parts, although here the CO2 treatments were effective only after bud set (Fig. 6b). The increase in total seedling biomass has also been observed in other commercial tree species. For instance, Sionit et al. (1985) reported that total dry weight increased by 56% in loblolly pine and 43% in sweetgum at 500 gL•L -1 CO2 compared with 350 gL•L -1 CO2. Working with black spruce, Campagna and Margolis (1989) indicated that total seedling biomass was 30 and 14% greater at 925 and 1100 11L•L -1 CO2, respectively, than control seedlings that did not receive CO2 enrichment. However, these authors found no stimulation of growth after bud set. Such differences are not surprising because it is known that the degree of response to CE can differ with respect to species, duration  66  of exposure, and stage of development (Brown and Higginbotham 1986, Kramer 1981, Sionit and Kramer 1986, Sionit et al. 1985, Tolley and Strain 1984). Looking at Fig. 6b, it can be noted that high CO2 treatment yielded the highest total seedling biomass at all stages of development, even where there were no statistical differences among treatments, at any given date. This suggests that, overall, CE enhanced dry matter production of Engelmann spruce seedlings, and that the effects were simply more apparent after bud set. The shoot:root ratio of Engelmann spruce seedlings in the present study (Fig. 8) compares favourably with that reported for other species (Campagna and Margolis 1989, Tolley and Strain 1984). In most CE studies with tree species, shoot:root ratio usually remains unchanged or slowly decreases as development progresses (Sionit et al. 1985). The present observations, however, show a sharp decline in shoot:root ratio at the end of the growing season (202 days in Fig. 7), although the treatment means were not statistically different. This decline in shoot:root ratio may partly be explained by the fact that shoot height growth ceases after bud set, whereas, root growth continues until limited by low temperature. Also the fact that roots were rapidly filling up with reserve carbohydrates could account for much of the increase in root dry weight.  67 5.2 Root Collar Diameter and Stem Height  Increases in stem diameter in response to CE have previously been demonstrated in sweetgum and loblolly pine (Sionit et al. 1985), ponderosa pine seedlings (Surano et a/. 1986), and in sweetgum and loblolly pine seedlings under CO2-enriched air with high irradiance (Tolley and Strain 1984). However, as already pointed out, such responses to CE are species specific, with some species responding more positively than others. The results of this study (Fig. 8a) also depicted similar trends in increased root collar diameter in response to CE. In this study, CE only significantly affected stem height of Engelmann spruce seedlings at 140 days (Fig. 8b). However, the general trend was that the high CO2 treatment attained greater height growth than the low CO2 treatment at all stages of development. The present observations are in agreement with the findings of Campagna and Margolis (1989) using black spruce, Surano et al. (1986) using ponderosa pine, and Sionit et al. (1984) using sweetgum and loblolly pine seedlings. It appears therefore, that for most species studied (the present study included), the effects of CE on growth are greater on biomass production than on stem diameter and stem height.  68  5.3 Carbohydrate Reserves  There is a broad body of literature indicating that carbohydrates (mostly starch and sugars) are the major form of food reserves for most woody plants (Duryea and McClain 1984, Glerum 1980a, 1980b, Kramer and Kozlowski 1979, Loescher et al. 1990, Ritchie 1984, 1987). In conifers, lipids and proteins are also important reserve materials (Glerum 1980a, Kramer and Kozlowski 1979), but there has been little work to establish just how important they are in relation to carbohydrates (Glerum 1980b). In temperate climates, reserve carbohydrates are a source of substrates for respiration during storage, and for early respiration, growth and development occuring in the subsequent year (Loescher et al. 1990, Marshall 1985, Ronco 1973). In conifers, reserve accumulation usually occurs in late summer and autumn (Gholz and Cropper, Jr. 1991, Glerum 1980b, Loescher et a/. 1990), and all plant parts function as sites of reserve accumulation and storage, but not with equal importance simultaneously (Glerum 1980b, Kramer and Kozlowski 1979). Generally speaking,^the trends^in^levels^of carbohydrate reserves observed in this study for Engelmann spruce seedlings are relatively similar to the ones reported for other species. In the present study, there was a general tendency for carbohydrate levels to decrease initially at 120 days, presumably due to the reduction in photoperiod (Figs. 9-16). The shortened photoperiod might have caused  69  the reduction in reserve carbohydrates because when photoperiod was reduced, there would have been fewer hours and hence less total light available for daily photosynthesis. In the case of TNC, CE was only effective after bud set, and by the end of the growing season (202 days in Fig. 9), the treatment effects were no longer apparent. The high TNC content seen at 202 days was mainly attributable to correspondingly large increases in starch in the roots and soluble sugars in the needles (which will be elaborated later). Carbon dioxide enrichment appeared to promote the rate of TNC accumulation following bud set, and this effect was apparent in both the high and step-up CO2 treatments. This effect was also seen in both major reserve pools (i.e. free sugars in the needles and starch in the roots). If high and step-up CO2 treatments can promote TNC accumulation after bud set, then it may be logical to provide CE after bud set instead of limiting its application to only the seedling emergence stage. Furthermore, if reserve carbohydrates in any way promote early growth and establishment of stored seedlings, and if these reserves are depleted during storage, then it is sensible to assume that any way in which TNC can be increased prior to storage might improve regeneration success. As indicated by this study, late season pre-storage CE might under some circumstances bring about some improvement in the whole plant TNC. The findings of this study differ from those reported by  70  Campagna and Margolis (1989), in that CE did not statistically alter TNC in black spruce seedlings. Again, these differences might simply reflect species specific response (Sionit et al. 1985, Tolley and Strain 1984, Sionit and Kramer 1986). Another possible explanation for the lack of an effect on TNC in black spruce could be that the duration of the CE treatment after bud set was insufficient. In terms of reserve allocation within the plant, roots constituted about 50% of the whole plant TNC just before storage (202 days), whereas, at all other times, needles had the higher proportion (Fig. 10). This suggests that, by the time seedlings were going into cold-storage, roots were the main storage organ. These results support the general observations and views by Abod and Webster (1991), Gholz and Cropper, Jr. (1991), Glerum (1980b), Loescher et a/. (1990), and McCracken (1979) that the root system is the main storage organ for total carbohydrate reserves in most tree species. The results of this study also conform to the findings of Glerum (1980b) and of Loescher et al. (1990) that the relative importance of different plant parts for storage of reserves varies at different stages of development.  71  Similarly, significant differences were observed in soluble sugar and starch content in response to CE by needles, stems and roots; particularly after bud set (Figs. 11-16). Campagna and Margolis (1989) observed similar effects in black spruce seedlings, but only prior to bud set. For soluble sugar content, needles recorded the highest values at all stages of development (Figs. 11-13). Soluble sugar contents in the stems and roots were comparable, although roots had slightly superior values. Here also, the findings of this study follow the trends in soluble sugar content reported elsewhere for other species (Abod and Webster 1991, Gholz and Cropper, Jr. 1991, Glerum 1980b, Kramer and Kozlowski 1979, Krueger and Trappe 1967). Starch content in all the three plant parts studied was affected by CE only at 140 days (Figs. 14-16), but starch accumulation and storage did not take place equally in different plant parts at different stages of development. Instead, roots were the major storage organ for starch, followed by stems, and in both cases starch accumulation rose sharply after bud set, reaching the maximum value achieved prior to cold-storage (202 days in Figs. 15 and 16). The very high build-up of starch in the roots and stems after bud set may reflect relative changes in the production and use of carbohydrates. Production would be expected to exceed use during this period since there is minimal growth, both above and below ground, while photosynthesis remains positive until the end of the season (unless set back by  72  repeated frost). Hagem (1941), and Krueger and Trappe (1967) speculated that, as diameter growth stops and root activity slows down in late October, starch and sugar reserves begin to increase gradually because, as autumn progresses, cool temperatures might reduce respiration. Furthermore, Alvik (1941) working with spruce and pine seedlings demonstrated that the average daylight and prevailing winter temperatures are usually sufficient to give a positive balance of assimilation over respiration for the greater part of the winter. Results presented here, where temperature was not reduced until 180 days, and yet carbohydrate reserves increased steadily, indicate that low temperatures may not be necessary to achieve positive carbon balance. The trends observed in Engelmann spruce seedlings follow closely those reported by Abod and Webster (1991), Glerum (1980b), Krueger and Trappe (1967), Loescher et al. (1990), and Reid et al. (1988). Based on the observations of this study, it appears that starch was the principal form of carbohydrate in the roots, whilst soluble sugars were more prevalent in the shoot, particularly in the needles. 5.4 Effects of Cold-storage on Carbohydrate Reserves  Loss of carbohydrate reserves during cold-storage has been widely studied in many temperate conifer species (Duryea and McClain 1984, Glerum 1980b, McCracken 1979, Ritchie 1982, 1984, 1987, Ronco 1972, 1973). These losses have been implicated as being responsible for poor growth  73  and establishment of outplanted seedlings. Therefore, one justification for storing seedlings at low temperature between lifting and planting is to minimize respiratory loss of carbohydrate reserves (Cleary and Tinus 1980 cited by Abod and Webster 1991). According to Ritchie (1987), coldstorage affects photosynthesis and respiration in two ways. First, the absence of light stops photosynthesis and second, low temperature decreases the rate of respiration. The net effect is that seedlings consume their supply of reserve carbohydrates in storage, but they do so very slowly. As stated by Duryea and McCracken (1984), Glerum (1980a), and Kozlowski and Kramer (1979), the primary use of carbohydrate reserves is to maintain respiration and growth when current photosynthate is not available. In this study, soluble sugars, starch and TNC were all affected by the two storage durations, but to varying degrees. On average, all the CO2 treatments behaved the same during the two storage durations. No effects were expected given the fact that all the CO2 treatments had almost the same TNC level when going into storage. For TNC, there was almost a one third reduction after two and four months (at 268 and 327 days in Fig. 9, respectively) in storage. Similar trends in TNC depletion have been observed in Engelmann spruce (Ronco 1972, 1973), in Mugo pine and radiata pine (McCracken 1979), in Douglas-fir (Ritchie 1982, 1987), and loblolly pine (Reid et a/. 1988) seedlings. However, it is interesting to note that, in this study,  74  there was a much greater apparent depletion of carbohydrate reserves between zero and two months storage than between two and four months storage. One possible reason for such a tendency in this study is that, when the cold-room temperature rose to about 10 ° C for two days at 230 days, seedlings might have respired excessively, thereby utilizing a larger portion of their reserves. However, it is likely that more reserve depletion occurred during the one week thawing period (at 5 ° C) just before planting. There was no such thawing period at 0 months storage as there was no need for one. In a study with spruce and pine seedlings, Alvik (1941) reported that respiration increased sharply between 0 ° and 10 ° C, and attained a temperature quotient (Q10) of 23. A change in Q 10 much greater than this would be required to account for the depletion observed in the present study. Still a temperature rise from -5 ° C (during storage) to 5 ° C (during thawing) might have increased the respiration rate of the seedlings substantially. The suspicion therefore, that thawing may contribute to the depletion of reserve carbohydrates, may suggest the need to examine the length and temperature of the thawing period for effects on the carbohydrate status of cold-stored seedlings. Changes in the Q10 for respiration at low temperature should be precisely established. Comparing plant parts, by the end of the two storage durations, roots had the highest reduction (more than 50%) in TNC and at these times, needles constituted more TNC than  75  any other plant tissue (Fig. 10). This is in agreement with other investigators who found similar reduction in root TNC after cold-storage (Philipson 1988, McCracken 1979, Ritchie 1982). On the whole, it appears that stem TNC was not affected by the two storage durations, and that stem TNC remained fairly constant throughout the whole study period (Fig. 10). Starch content within plant tissues was significantly affected by the two and four months storage durations, and the depletion of starch during the two storage durations was comparable (at 268 and 327 days in Figs. 14-16). In all the three plant parts studied, there was more than 50% starch depletion as compared with the starch levels prior to coldstorage (i.e. at 202 days). While the results of this study conform to those of Ronco (1972, 1973) for Engelmann spruce, and McCracken (1979) for Mugo and radiata pines, conflicting results were reported by Krueger and Trappe (1967), and Ritchie (1982) for Douglas-fir seedlings. Both of these studies indicated an increase in starch content between March and April (Krueger and Trappe 1967), and after nine months storage (Ritchie 1982). As regards soluble sugars, this study has shown that the two and four months storage durations did not have much impact on their depletion (at 268 and 327 days in Figs. 1113). There was only about 5 and 25% reduction in needle and root soluble sugars, respectively. In studies with Engelmann spruce (Ronco 1972, 1973), Douglas-fir (Ritchie 1982,  76  Krueger and Trappe 1967), and Mugo and radiata pines (McCracken 1979), the pattern in the depletion of soluble sugar content in needles, stems, and roots was essentially identical to the results of the present study. The only difference was that, in this study, soluble sugar content seems to have been maintained at the expense of starch, whereas, in the former experiments soluble sugar contents were not static and declined almost linearly with storage duration. Furthermore, since respiration presumably occurred throughout the plant and seedlings were not photosynthesizing while in storage, it is reasonable to assume that the maintenance of needle sugar content was due to starch conversion in roots to soluble sugars, which were then translocated to the needles. This observation suggests that, at least in Engelmann spruce, efficient phloem transport between the roots and needles may occur in storage. The enzymatic reaction involved in maintaining the equilibrium between starch and free sugars is somewhat temperature dependent. Low temperature favours conversion of starch to free sugars (ap Rees et al. 1988, DeLucia 1986, Ritchie 1982), but the actual control of this process is not well understood. Specific experiments should be designed to examine circumstances surrounding starch conversion and phloem transport in Engelmann spruce. On the whole, the results from this study strongly  conform to the already established fact that carbohydrate levels of cold-stored seedlings deplete with storage  77  duration. However, there are some differences in the level of depletion depending on the individual species, lifting date, storage temperature, storage duration, and other cultural practices (Duryea and McClain 1984). This study also indicates that pre-storage CE per se did not have any influence on the degree of TNC, starch and soluble sugar depletion in Engelmann spruce seedlings while in storage. 5.5 Bud Break  The data reported in this study indicate that none of the CO 2 treatments showed any significant influence on the days to first bud break at either 268 or 327 days (Table 1). However, at 268 days, where soil temperature was considered as a factor, bud break was significantly affected by soil temperature. It is important to point out here that, although soil temperatures differed statistically, the mean days to first bud break did not differ greatly, especially between 7 and 11 ° C soil temperatures. These results support the findings of Cam and Harper (1991) who reported similar trends in days to terminal bud break in white spruce seedlings after 15 weeks of cold-storage. Results from soil temperature studies on Douglas-fir, Pacific silver fir (Abies amabilis (Dougl.) Forbes), noble fir (Abies procera Rehd), lodgepole pine, and ponderosa pine seedlings (Lopushinsky and Max 1990) also showed decreasing days to bud break with increasing soil temperature. However, even with as wide as 0-30 ° C soil temperature range the true  78  firs did not respond as strongly to the increasing soil temperature as did Douglas-fir and the pines. Earlier work by Lavender and Overton (1972) reported that the reduction in shoot growth of Douglas-fir seedlings associated with low soil temperature was not occasioned by reduced water or mineral uptake. Their alternative explanation was that when roots are grown in cold soils, shoot growth or bud activity is slowed by reduced export, from the roots, of some plant growth regulatory substance or substances (presumably gibberellins). Similar conclusions were reached by Lavender and Wareing (1972), and Lavender et al. (1973) who found that in Douglas-fir transplants, gibberellin-like compounds were synthesized in the roots and then exported to the shoots where they enhanced bud activity. The decrease in days to first bud break with increasing cold-storage duration is also in conformity with what other researchers have reported. For example, Camm and Harper (1991) working with white spruce found a similar pattern in days to bud break over a wide range of cold-storage durations (0-30 weeks). Similarly, several authors (Burr et al. 1989, Carlson 1985, Ritchie 1984, Ritchie et al. 1985, van den Driessche 1977) working with a wide range of species also reported decreasing days to terminal bud break with increasing storage duration. This decrease in days to bud break may reflect the accumulation of chilling hours required to fully release dormancy. Chilling requirements in spruce are usually satisfied after 6-8 weeks of exposure to  79  low temperatures (ca. 4-6 ° C) (Lavender unpublshed data, Nienstaedt 1966, 1967). As noted by many researchers (Burr et al. 1989, Carlson 1985, Lavender 1985, Ritchie and Dunlop 1980, Ritchie et al. 1985), accumulation of chilling hours required to release dormancy for most temperate conifer species can be achieved by placing seedlings in storage after they have entered rest, usually in late November or early December. It is also known that chilling interruption by brief periods of warmer temperature during storage can affect dormancy release, and that the degree of negating the chilling response depends on timing, duration, and temperature during the interruption period (Lavender: personal communication, van den Driessche: cited by Ritchie 1984,). In this study, a brief interruption of cold-storage at 230 days (i.e. 10 ° C for two days) may have influenced days to first bud break. Early bud break may be beneficial at locations with short growing periods (e.g. interior B.C.), in that, outplanted seedlings can resume growth early to take advantage of the short favourable growing season. Equally important, a delay in dormancy release during cold-storage can be used to the advantage of a practising forester in significantly expanding the planting "window". However, one must be cautious to ensure that any storage duration or protocol chosen does not significantly promote depletion of carbohydrate reserves. As already indicated in this study, and by numerous researchers, prolonged storage durations  80  negatively affect carbohydrate levels in most forest tree seedlings. It has not, however, been well-established, here or elsewhere, as to what degree of reserve depletion is tolerable. Substantial, almost complete loss of reserves may be inconsequential (Marshall 1985, Omi and Rose 1990, Ritchie 1982, 1984, Ronco 1973), but complete exhaustion of reserves is very likely fatal. 5.6 Root Growth Potential  Although it has long been suspected that carbohydrate reserves are important for outplanted seedlings after storage, the direct relationships between reserves and early root growth after cold-storage have not been thoroughly explored. There are still numerous conflicting results as to whether new root growth after cold-storage depends on "old" carbon (stored carbohydrates) or "new" carbon (current photosynthate) or both.  5.6.1 Effects of CE and Soil Temperature on RGP  In the present study, none of the CO2 treatments had any positive effects on RGP of Engelmann spruce seedlings at both 268 and 327 days (Table 2). Since all the CO2 treatments had almost the same TNC level prior to coldstorage, it was not surprising, at least in terms of reserve utilization, that RGP was not affected by CE after storage. On the other hand, the significant effect observed at 268 days in response to varying soil temperatures is in  81  agreement with the results reported by many investigators. In a study with white spruce, Camm and Harper (1991) indicated that, after 17.5 weeks of cold-storage, initiation of new root growth was soil temperature-dependent. The results reported in this study compare favourably with those of Camm and Harper (1991) for white spruce, and are above the RGP threshold value for adequate root regeneration (10 new roots > 10mm long) for interior spruce and lodgepole pine reported by Simpson et al. (1988). Reduced root growth at low soil temperatures is not surprising because similar findings have been reported for other conifers. In an experiment with Douglas-fir, Pacific silver fir, noble fir, lodgepole pine, and ponderosa pine transplants, Lopushinsky and Max (1990) indicated that maximum root growth in all species occurred at 20 ° C. Furthermore, soil temperatures of 10 ° C or less drastically reduced or prevented new root growth. Other studies with Douglas-fir seedlings (Lopushinsky and Kaufmann 1984, Ritchie 1985) also showed that little root growth occurred at soil temperatures of 5 ° C or less. In contrast, Lavender and Wareing (1972) reported improved root growth in Douglas-fir seedlings at 4 ° C soil temperature as a result of application of gibberellin-like compounds. If rapid initiation of new root growth is essential for the early establishment of outplanted seedlings, and if new root growth does not generally proceed well at soil temperatures below 10 ° C, it is logical to suggest that seedlings should not be planted until soils  82 warm up to 10 ° C or more. However, in many situations, waiting for the soil temperature to warm up to 10 ° C. would mean extending storage duration which in turn may affect the seedling quality for a variety of reasons, including reserve depletion. Another possibility would be to move towards summer lifting/and planting (i.e. so-called "hot" planting). The development of RGP in conifer species tends to follow a general pattern; low in fall, reaching a peak in mid-winter, declining in spring, and then rising slightly again in late summer (Ritchie 1985, Ritchie and Dunlop 1980, Ritchie and Tanaka 1990, Stone and Jenkinson 1970). In addition to soil temperature and physiological condition (particularly dormancy status), cold-storage in interaction with lifting date may also have a strong effect on the expression of RGP (Ritchie 1985). Generally speaking, seedlings lifted in mid-winter tend to have higher RGP than fall or spring-lifted seedlings (Burdett 1987, Ritchie and Dunlop 1980, Ritchie et al. 1985, Silim and Lavender 1992, Sutton 1990). They also have higher RGP following storage for a few months.  5.6.2 Effects of Cold-storage on RGP  In this study, there was no clear pattern in the RGP of Engelmann spruce seedlings with different storage duration as assayed at a soil temperature of 11 ° C (Table 2). The results from this study therefore are not consistent with the general contention that, for most conifer species, RGP  83  increases during the initial stages of cold-storage, and then declines with prolonged cold-storage duration. On the whole, RGP values are notoriously variable, and trends observed between and even within different studies are often inconsistent, partly due to differing test environments, management practices, and methods of evaluating RGP (Ritchie and Dunlop 1980, Ritchie et a/. 1990, Sutton 1990). For instance, Carlson (1985) working with loblolly pine seedlings found that RGP in fall-lifted seedlings was reduced during cold-storage, whereas storage after midwinter (after 734 chilling hours) either improved or did not affect RGP. Ritchie et a/. (1985) studied RGP in lodgepole pine and interior spruce seedlings, and in both species, December to March lifting dates, followed by a two month storage had little effect on RGP. However, when the same seedlings were subjected to a six month storage, RGP was substantially reduced especially for the March lift date. In separate experiments, Camm and Harper (1991) and Ritchie (1982), working with white spruce and Douglas-fir seedlings respectively, observed that RGP steadily increased during the initial stages of cold-storage (ca. 0-4 months), but declined sharply after prolonged storage. The physiological condition of the seedlings at the initiation of storage could be viewed as a major factor controlling RGP periodicity. For example, Ritchie and Dunlop (1980) speculated that seasonal patterns in carbohydrate synthesis, storage, conversion, and metabolism may be the  84  primary modulators of RGP periodicity in trees. They further stated that the highest concentration of carbohydrates during mid-winter happens to coincide with the presumed RGP and stress resistance peaks, after which RGP declines as carbohydrate reserves deplete during storage. This coincidence between carbohydrate levels and RGP may have encouraged the concept that carbohydrate reserves are important substrates for root growth.  5.5.3 Role of Carbohydrates in RGP  Several studies on the role of carbohydrate reserves in RGP have presented conflicting results. However, the dependence of outplanted seedlings on either carbohydrate reserves, current photosynthate or a combination appears to be somewhat species specific. The results from this study indicate that after 9, 18, and 36 days, the new roots of planted seedlings of Engelmann spruce seedlings were respectively, comprised of 100, 53, and 18% "old" carbon (Table 4). This suggests that, at least during the first 9 days, new root growth of planted Engelmann spruce seedlings depends entirely on carbohydrate reserves; although there was very little root growth during this initial period. There is no doubt, however, that reserve carbon did contribute significantly to root construction up to 36 days after planting. These observations support the contention of many authors that new root growth in some species depends, at least in part, on carbohydrate reserves. For instance, in  85  disbudding and girdling or bark-ringing experiments, Philipson (1988) reported that new spring root growth of Sitka spruce transplants depended on carbohydrates stored within the root. By way of radioisotope experiments, van den Driessche (1987) obtained contradictory results; new root growth for both Douglas-fir and Sitka spruce seedlings seemed to depend on current photosynthate. Lavender and Hermann (1970), and Lavender and Wareing (1972) also reported that new root growth of Douglas-fir transplants may largely depend on some substance or substances, presumably hormones or carbohydrates, exported from the foliage. These findings match with those of Ursino et al. (1968), who also reported that the new roots of white pine (Pinus strobus L.) transplants received current photosynthate from the shoot throughout the entire growing season. However, neither girdling nor radioisotope experiments can provide unequivocal results useful for finally resolving the controversy. For example, bark-ringing experiments may not only exclude translocation of current photosynthate down to the root system, but other known or unknown compounds as well. As noted by Philipson (1988), although new root growth in Sitka spruce seedlings was not dependent on current photosynthate, after 14 days, bark-ringed trees started to show retarded growth, suggesting that the roots were lacking some factor. The main shortfall in using radioisotopes for C-labelling is that the labelling period is short (ranging from one hour to a couple of days) and, as such, may not  86  provide a uniform or thorough labelling of reserves. Stable carbon isotope labelling, as used in this study should provide a better technique for labelling and then tracing the allocation of "old" carbon to a given plant part (new roots in this case). In girdling experiments it is very difficult to implicate lack of current photosynthate as the sole factor inhibiting new root growth. It has been speculated by many researchers that certain unknown compounds (presumably plant growth regulators), which are translocated along with current photosynthate to the roots also play some role in new root growth (Deyoe and Zaerr 1976, Kramer and Kozlowski 1979, Philipson 1988, Ritchie and Dunlop 1980, Zaerr and Lavender 1974). Van den Driessche (1991) investigated the effect of low CO2 concentration, darkness, and stem girdling on new root growth in Douglas-fir seedlings. He found that new root production was affected by the treatments in the order: control > low CO2 concentration > dark > girdling. Production of new roots was limited in the absence of photosynthesis (i.e. dark treatment) but was even more seriously affected by girdling, suggesting that lack of current photosynthate and presumably other factors might have inhibited new root growth. His findings supported the views of Lavender and Hermann (1970), Philipson (1988), Ritchie (1982), van den Driessche (1987), and Zaerr and Lavender (1974) that darkening or girdling almost completely prevents new root production. Girdling treatment was more  87  severe in inhibiting new root production presumably because only reserves stored in the root system would be accessible, whereas, in the low CO2 concentration and dark treatments, reserves throughout the whole plant would have been accessible.  88  6.0 CONCLUSIONS  The principal objective of this thesis research was to establish the role of carbohydrate reserves in spring root growth of freezer-stored Engelmann spruce seedlings. Emphasis was placed on investigating the utility and possible side effects of CE in the manipulation of these reserves. Based on the foregoing results and discussion, the following salient conclusions can be drawn from the present study:  1. Biomass production by individual plant parts in Engelmann spruce seedlings was more significantly affected than root collar diameter and stem height by CE. These effects were more apparent after bud set.  2. Despite the fact that biomass production of individual plant parts was affected by CE, shoot:root ratio was not affected by CE, and gradually decreased during the study period (i.e. 80-202 days).  3. Carbon dioxide enrichment had little significant effect on carbohydrate levels prior to bud set.  4. In contrast to the above, CE did promote the rate of carbohydrate accumulation following bud set, but had no influence on the ultimate level attained just before storage.  89  5. After two and four months storage, there was almost a one-third reduction in the whole plant TNC.  6. Soluble sugars in all plant parts were not significantly affected by either two or four months storage whereas, there was more than 50% reduction in starch in all plant parts following two and four months storage.  7. Carbon dioxide enrichment did not have any detectable effect on days to bud break or RGP.  8. Low soil temperature increased the number of days to bud break and adversely affected RGP.  9. Stored reserves made an important contribution to early spring root growth, but this sink was only a minor drain on the reserve pool.  90  7.0 RECOMMENDATIONS  The results from this study may be of some practical use in the reforestation program in B.C. (especially northerly areas) in the following ways:  1.  In most of the parameters measured (i.e. biomass  production, growth variables, and carbohydrates), the effects of CE were most apparent after bud set. Therefore, nursery operators should consider CE of Engelmann spruce seedlings applied after bud set to benefit growth and manipulate seedling morphology. Carbon dioxide enrichment following bud set would effectively increase biomass independently of shoot height growth.  2.  Extended storage and/or thawing periods should be  avoided as much as possible. As noted by Marshall (1985), and Omi and Rose (1990) the degree of carbohydrate depletion during storage will depend on the amount of reserves going into storage and subsequent storage duration and temperature. There is some potential for the use of CO2 enrichment for enhancement of reserve levels prior to storage.  91  3 Future Research  Owing to time constraints and the fact that factors affecting root regeneration, especially in cold soils are diverse, it was not feasible to tackle all aspects related to root regeneration in this one thesis project. Therefore, the following are some areas in need of future research:  - The effects of thawing period and temperature on the depletion of carbohydrates following cold-storage, should be investigated.  - The storage period should be extended to six months as initially planned in the present study to cover a more complete range of storage durations that might demonstrate adverse effects of prolonged cold-storage in terms of carbohydrate depletion and expression of RGP.  - The observed effects of CE after bud set should be investigated for possible impacts on seedling field performance.  - The role of roots as a major site of storage suggests that more emphasis should be placed on understanding root hardiness, mobilization of root reserves, etc.  92  - The CE experiment should be repeated. This should be done after bud set for a shorter duration (ca. 6-7 weeks). Also a greater range of CO2 concentrations should be tested.  -  In repeating the stable carbon isotope labelling  experiment, the harvesting dates should be extended from 36 days to a point where it can be demonstrated that all the reserve carbon in the roots of planted seedlings is exhausted.  - In repeating the stable carbon isotope labelling experiment, per cent utilization of reserves by other sinks (e.g. buds) should be established.  93 8.0 LITERATURE CITED  Abod, S.A., and A.D. Webster. 1991. Carbohydrates and their effects on growth and establishment of Tilia and Betula: I. Seasonal changes in soluble and insoluble carbohydrates. J HortSci 66:235-246. Alvik, G. 1941. On assimilation and respiration in some woody plants under western-Norwegian winter conditions. For Abstr 2:189-90. ap Rees, T., M.M. Burrell, T.G. Entwistle, J.B.W. Hammond, D. Kirk, and N.J. Kruger. 1988. Effects of low temperature on the respiratory metabolism of carbohydrates by plants. In: S.P., and F.I. 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Prentice Hall, Englewood Cliffs, NJ. 718 p.  103 Appendix 1 ANOVA for the effect of CE on needle, stem, and root dry weights in Engelmann spruce seedlings.  (A) Needle dry weight at 80 days. ^  ^  ^  F Ratio P DF^MS ^ Treatments^73154.533 ^ 3^24384.844^3.010^0.034 92^8102.047 Error^745388.366 SV  SS  -  (B) Stem dry weight at 80 days. SV  ^  SS^DF  ^  MS  ^  F-Ratio P  ^ ^ Treatments ^ 8892.862 ^ 3^2964.287^4.683^0.004 Error 58231.422 92^632.950 (C) Root dry weight at 80 days. ^  ^  ^  F-Ratio DF^MS ^ ^ Treatments ^ 31948.815^ 3^10649.605^6.853^0.000 Error 142964.422 92^1553.961 SV  SS  (D) Needle dry weight at 120 days. SV  ^  SS  ^  DF^MS  ^  F-Ratio  Treatments 244066.294^3^81355.431^1.958^0.126 Error^3823036.971^92^41554.750 (E) Stem dry weight at 120 days. ^  ^  ^  F-Ratio DF^MS ^ ^ ^ 2.083^0.108 Treatments 180.495 ^ 3^14393.498 ^ Error 635763.184 92^6910.469 SV  SS  104 (F) Root dry weight at 120 days. SV^SS^DF^MS  F Ratio -  1.688  0.175  SV^SS^DF^MS  F Ratio  P  Treatments^680709.781 Error^4549861.254  11.328  0.000  Treatments^69206.005^3^23068.668 Error^1257376.160^92^13667.132 (G) Needle dry weight at 140 days.  3^560236.594 92^49455.01  -  (H) Stem dry weight at 140 days. SV^SS  DF^MS  Treatments^387515.884 Error^1210320.649  3^129171.961 92^13155.659  F Ratio -  9.819  0.000  (I) Root dry weight at 140 days. SV^SS  DF^MS  F Ratio  Treatments^837655.114 Error^2373570.919  3^279218.371 92^25799.684  10.823  -  0.000  (J) Needle dry weight at 202 days. SV^SS  DF^MS  F Ratio  Treatments^734801.893 Error^695197.634  3^244933.964 24^28966.568  8.456  0.001  F Ratio  P  -  (K) Stem dry weight at 202 days. SV^SS  DF^MS  Treatments^167519.070 Error^273318.020  3^55839.690 24^11388.251  -  4.903  0.008  (L)^Root dry weight at 202 days. SV^SS  DF^MS  Treatments^449830.473 Error^1339381.737  3^149943.491 24^55807.572  F Ratio -  2.687  0.069  105 Appendix 2^ANOVA for the effect of CE on total biomass production in Engelmann spruce seedlings.  (A) Total seedling biomass at 80 days. SV^SS^DF^MS^F-Ratio^P Treatments^277816.235^3^92605.412^4.620^0.005 Error^1843997.912 92^20043.456  (B) Total seedling biomass at 120 days. SV  ^  SS^DF^MS^F-Ratio^P  Treatments 810464.168 ^3^270154.723^1.980^0.122 Error^.125533E+08^92^136449.202  (C) Total seedling biomass at 140 days. SV^SS^DF^MS^F-Ratio^P Treatments 8062351.407^3 2687450.469^12.478^0.000 Error^.198144E+08^92^215373.882  (D) Total seedling biomass at 202 days. SV^SS^DF^MS^F-Ratio^P Treatments 3351344.275 ^3 1117114.758^8.258^0.001 Error^3246539.314^24^135272.471  106 Appendix 3 ANOVA for the effect of CE on shoot:root  ratios in Engelmann spruce seedlings.  (A) Shoot:root ratios at 80 days. SV^SS^DF^MS^F-Ratio Treatments 5.001^3^1.667^2.512^0.063 Error^61.064^92^0.664  (B) Shoot:root ratios at 120 days. SV  ^  SS^DF^MS^F-Ratio  Treatments^9.270^3^3.090^6.478^0.001 Error^43.887 92^0.477  (C) Shoot:root ratios at 140 days. SV  ^  SS^DF^MS^F-Ratio  Treatments^3.401^3^1.134^1.884^0.138 Error^55.356 92^0.602  (D) Shoot:root ratios at 202 days. ^  SS^DF^MS^F-Ratio ^ ^ 1.610^0.209 Treatments 1.538^3 ^0.513 Error^8.917 28 0.318 SV  107  Appendix 4 ANOVA for the effect of CE on diameter and stem height growth in Engelmann spruce seedlings.  (A) Diameter growth at 80 days. SV  ^  SS^DF^MS^F-Ratio  Treatments 0.418^3^0.139^4.469^0.006 Error^2.869^92^0.031 (B) Diameter growth at 120 days. SV  ^  SS^DF^MS^F-Ratio  Treatments 0.534^3^0.178^1.531^0.212 Error^10.684^92^0.116 (C) Diameter growth at 140 days. SV  ^  SS^DF^MS^F-Ratio  Treatments 4.173^3^1.391^9.334^0.000 Error^13.710^92^0.149 (D) Diameter growth at 202 days. SV  ^  SS^DF^MS^F-Ratio  Treatments 0.564^3^0.188^5.028^0.008 Error^0.898^24^0.037 (E) Stem height growth at 80 days. SV  ^  SS^DF^MS^F-Ratio  Treatments 3.968^1^3.968^2.099^0.154 Error^86.943^46^1.890 (F) Stem height growth at 120 days. ^  SS^DF MS^F-Ratio ^ Treatments 12.403^1 12.403 1.538^0.221 Error^371.069^46^8.067 SV  108 (G) Stem height growth at 140 days. ^  SS^DF MS^F-Ratio^P ^ 12.003^0.001 Treatments 93.800^1^93.800 Error^359.490^46^7.815 SV  Appendix 5 ANOVA for the effect of CE on carbohydrate reserves at 80 days. (A) Starch in needles. SV^SS  DF  Treatments^14735.095 Error^108447.059  3 44  MS 4911.698 2464.706  F-Ratio  P  1.993  0.129  F-Ratio  P  2.143  0.108  F-Ratio  P  (B) Starch in stems. SV^SS  Treatments^747.122 Error^5114.314  DF 3 44  MS 249.041 116.234  (C)^Starch in roots. SV^SS  DF  MS  Treatments^941.146^3^313.715^1.174^0.331 Error^11759.898^44^267.270 (D) Soluble sugars in needles. SV^SS^DF^MS^F Ratio^P -  Treatments^36137.536^3^12045.845^1.463^0.238 Error^362206.300^44^8231.961 (E) Soluble sugars in stems. SV  ^  SS^DF^MS^F-Ratio^P  Treatments^38420.142^3^12806.714^9.088^0.000 Error^62002.888^44^1409.157  109 (F) Soluble sugars in roots. SV^SS^DF^MS^F-Ratio ^ 24.651^0.000 Treatments 104512.767^3^34837.589 Error^62181.038^44^1413.205 (G) Whole plant TNC. SV  ^  SS^DF^MS^F-Ratio  Treatments^6214.097^3^2071.366^0.478^0.699 Error^190637.767^44^4332.677  Appendix 6 ANOVA for the effect of CE on carbohydrate reserves at 120 days. (A) Starch in needles. SV^SS^DF^MS^F-Ratio^P Treatments^1670.615^3^556.872^2.242^0.097 Error^10929.628^44^248.401 (B) Starch in stems. SV^SS^DF^MS^F-Ratio Treatments^257.559^3^85.853^1.865^0.149 Error^2025.193^44^46.027 (C) Starch in roots. SV^SS^DF^MS^F-Ratio Treatments^211.456^3^70.485^0.626^0.602 Error^4952.226^44^112.551 (D) Soluble sugars in needles. SV  ^  SS^DF MS^F-RATIO  Treatments^35034.436^3 11678.145^4.097^0.012 Error^125431.529^44^2850.717  110  (E) Soluble sugars in stems. SV^SS^DF^MS^F-RATIO^P Treatments^20985.172^3^6995.057^3.842^0.016 Error^80108.979^44^1820.659 (F) Soluble sugars in roots. SV^SS^DF^MS^F-RATIO Treatments^8039.302^3^2679.767^5.509^0.003 Error^21402.470^44^486.420 (G) Whole plant TNC. SV^SS^DF^MS^F-Ratio Treatments^21564.213^3^7188.071^5.571^0.002 Error^56770.012^44^1290.228  Appendix 7 ANOVA for the effect of CE on carbohydrate reserves at 140 days. (A) Starch in needles. SV^SS^DF^MS^F-Ratio Treatments^17699.206^3^5899.735^4.931^0.005 Error^52647.756^44^1196.540 (B) Starch in stems. SV^SS^DF^MS^F-Ratio  Treatments^13241.021^3^4413.674^6.916^0.001 Error^28080.447^44^638.192 (C) Starch in rootss. SV^SS^DF^MS^F-Ratio Treatments^89677.812^3^29892.604^9.478^0.000 Error^138766.489^44^3153.784  111  (D) Soluble sugars in needles. SV^SS^DF^MS^F-Ratio^P Treatments 189912.464^3^63304.155^14.727^0.000 Error^189135.203^44^4298.527 (E) Soluble sugars in stems. SV^SS^DF^MS^F-Ratio Treatments^19322.194^3^6440.731^2.707^0.057 Error^104680.250^44^2379.097 (F) Soluble sugars in roots. SV^SS^DF^MS^F-Ratio Treatments^11484.508^3^3828.169^3.488^0.023 Error^48292.811^44^1097.564 (G) Whole plant TNC at 140 days. SV^SS^DF^MS^F-Ratio Treatments^171572.186^3^57190.729^19.590^0.000 Error^128451.948^44^2919.362  Appendix 8 ANOVA for the effect of CE on carbohydrate reserves at 202 days.  (A) Starch in needles. SV^SS  DF  Treatments^4264.294 Error^17165.508  3 25  MS 1421.431 686.620  F-Ratio 2.070  0.130  (B) Starch in stems. SV^SS Treatments^961.970 Error^40891.470  DF 3 25  MS 320.657 1635.659  F-Ratio 0.196  0.898  112 (C) Starch in roots. SV^SS^DF^MS^F-Ratio Treatments^4793.192^3^1597.731^0.261^0.853 Error^153042.129^25^6121.685 (D) Soluble sugars in needles. SV^SS^DF^MS^F-Ratio Treatments^27341.579^3^9113.860^3.280^0.037 Error^69460.530^25^2778.421 (E) Soluble sugars in stems. SV^SS^DF^MS^F-Ratio^P Treatments^2750.799^3^916.933^3.381^0.034 Error^6779.269^25^271.171 (F) Soluble sugars in roots. SV^SS^DF^MS^F-Ratio ^ 11.803^0.000 Treatments^32930.876^3^10976.959 Error^23250.298^25^930.012 (G) Whole plant TNC. SV^SS^DF^MS^F-Ratio ^ Treatments 10358.780^3^3452.927 2.109^0.125 Error^40935.990^25^1637.440  Appendix 9 ANOVA for the effect of CE on carbohydrate reserves at 268 days.  (A) Starch in needles. SV^SS^DF^MS^F-Ratio^P Treatments^10.131^3^3.377^3.163^0.064 Error^12.813^12^1.068  113 (B) Starch in stems. SV  SS^DF^MS^F-Ratio^P  Treatments 1877.587^3^625.862^0.818^0.508 Error^9178.559^12^764.880 (C) Starch in roots. SV  SS^DF^MS^F-Ratio^P  Treatments 8489.077^3^2829.692^0.771^0.532 Error^44061.791^12^3671.816 (D) Soluble sugars in needles. SV^SS^DF^MS^F-Ratio^P Treatments 6849.386^3^2283.129^2.131^0.150 Error^12856.853^12^1071.404 (E) Soluble sugars in stems. SV^SS^DF^MS^F-Ratio^P Treatments 2925.569^3^975.190^2.822^0.084 Error^4147.044^12^345.587 (F) Soluble sugars in roots. SV^SS^DF^MS^F-RATIO^P Treatments^13713.960^3^4571.320^3.547^0.048 Error^15467.311 12^1288.943 (G) Whole plant TNC. SV  SS^DF^MS^F Ratio^P -  Treatments^9915.705^3^3305.235^1.957^0.174 Error^20264.445^12^1688.704  114 Appendix 10 ANOVA for the effect of CE on carbohydrate  reserves at 327 days.  (A) Starch in needles. SV  ^  SS^DF^MS^F-Ratio  Treatments^8.870^3^2.957^4.024^0.022 Error^14.696^20^0.735 (B) Starch in stems. SV^SS^DF^MS^F-Ratio Treatments^760.760^3^253.587^0.175^0.912 Error^8910.440^20^1445.522 (C) Starch in roots. SV^SS^DF^MS^F-Ratio^P Treatments^3671.468^3^1223.823^0.670^0.580 Error^36507.081^20^1825.354 (D) Soluble sugars in needles. SV^SS^DF^MS^F-Ratio Treatments^5410.845^3^1803.615^4.960^0.010 Error^7272.851^20^363.643 (E) Soluble sugars in stems. SV^SS^DF^MS^F-Ratio Treatments^5710.163^3^1903.388^1.625^0.215 Error^23423.801^20^1171.190 (F) Soluble sugars in roots. SV^SS^DF^MS^F-Ratio Treatments^5421.281^3^1807.094^1.231^0.325 Error^29362.064^20^1468.103  115 (G) Whole plant TNC. ^ ^ F-Ratio DF MS ^ Treatments^3711.179 ^ 3^1237.060^1.580^0.225 20^782.747 Error^15654.940  SV^ SS  ^  Appendix 11 ANOVA for the effect of CE on bud break and RGP in Engelmann spruce seedlings.  (A) Bud break at 268 days. SV  ^  SS  ^  Water baths 24.942 Treatments 18.724 Water bath*treat. 14.162 Error 139.348  DF^MS^F-Ratio  2 5 10 96  8.592 2.580 0.976  12.471 3.745 1.416 1.452  0.000 0.031 0.470  (B) Bud break at 327 days. SV  DF  SS  Water baths 1.975 Treatments 95.740 Water bath*treat. 78.016 Sampling error 953.524  F-Ratio  MS  0.987 19.148 7.802 10.364  2 5 10 92  P 0.909 0.111 0.673  0.095 1.847 0.753  (C) RGP at 268 days. SV  DF  SS  Water baths 18113.389 Treatments 5798.611 Water bath*treat. 3419.111 Error 61310.667  1 5 5 60  F-Ratio  MS  18113.389 1159.722 683.822 1021.844  17.726 1.135 0.669  P 0.000 0.352 0.648  (D) RGP at 327 days. SV  ^  SS  ^  Water baths 4383.865 Treatments 17850.197 Water bath*treat. 14856.357 Sampling error 145437.357  DF 2 5 10 89  ^  MS  ^  2191.932 3570.039 1485.636 1634.128  F-Ratio 1.341 2.185 0.909  ^  P  0.267 0.063 0.528  116 Appendix 12. Protocol for Carbohydrate Analyses. I.  Grinding the plant material (needles, roots and stems)  Grind the freeze-dried plant material in liquid nitrogen using a mortar and pestle. Needles are easily crushed by this method, but roots and stems may be difficult to grind and may require milling. The ground material can be stored in vials at 2 - 6°C until required for analysis. Tissues ground in liquid nitrogen pick up some condensation, and should therefore be dried again prior to storage. II.  Separating soluble sugars from starch  1.  Weigh out 20 to 50 mg of the pulverized plant material and put it in a test tube.  2.  To each test tube add 5 mL of methanol: choloroform: water (M:C:W - 12:5:3, v/v/v) and leave the samples overnight.  3.  The following morning, remove the samples from the freezer, briefly shake them on a vortex and then centrifuge for 10 minutes (desk top). Pipet the supernatant containing soluble sugars into vials and save. Starch will remain with the pellet.  4.  Add 3 mL of M:C:W to the pellet in the test tube. Vortex briefly and then centrifuge for 10 minutes (desk top).  5.  Pipet off the supernatant and combine with the one from step 3.  6.  Store the pellet in the freezer or take it immediately for starch analysis.  III. Final extraction of soluble sugars  1.  To each vial of supernatant from above, add 3 mL of distilled water. Shake the vials gently by hand and let settle (about 5 minutes).  2.  Pipet the top aqueous layer into an evaporating flask. The lower layer is waste, mostly chloroform. Flash evaporate all the solvent from the flask, while the flask rotates in a 40°C water bath. It takes 5 - 10 minutes to do one evaporation.  3.^Add 1.5 mL of distilled water to the flask, swirl gently and pipet into a microcentrifuge tube. Wash the flask with another 1.5 mL of distilled water, swirl gently and add to the microcentrifuge tube. Spin for 3 minutes and transfer the supernatant into vials and store solutions in the freezer until required for sugar analysis.  117 IV. Starch analysis using enzymes  1.  To the pellet in the test tube, add 5 mL of acetate buffer (pH 4.5, 150 mM) and briefly vortex.  2.  Autoclave at 120°C for 1 hour or incubate in a water bath at 88°C for 3 hours.  3.  Put the samples in a 55°C water bath for about 5 - 10 minutes.  4.  Add to each test tube: - 62.5 pL a-amylase solution (see VI.3) - 125 gL amyloglucosidase solution (see VI.2) Gently shake the contents by hand to mix the enzymes and the starch solution. Incubate for 2 hours then cool for 5 - 10 minutes at room temperature.  5.  Centrifuge for 10 minutes (desk top).  6.  Aiming for a working volume of 200 AL, pipet from step 5 the desired amount of aliquot and dilute it appropriately. This requires some sense of what the starch concentration in a particular plant extract might be (i.e., varies with time of year, plant part, etc.). For spruce, the following are usually reasonable: Needles: 200 AL plant material + 0 AL acetate buffer Roots: 100 AL plant material + 100 pL acetate buffer Stems: 150 AL plant material + 50 AL acetate buffer.  7.  Add 2 mL of combined peroxidase/glucose oxidase/colour reagent solution.^See section VI.4 or refer to the attached Sigma Diagnostics Procedure No. 510.  8.  Incubate the samples for 30 minutes in a 37°C water bath.  9.^Read the absorbance at 450 nm. Refer to: HAISSIG, B.F. and DICKSON, R.E. 1979, Physiol Plant 47:151-157. Starch concentrations are calculated in glucose equivalents. When establishing a standard curve, first prepare a starch standard solution by dissolving 16.2 mg of soluble starch powder (BDH chemicals, ACS 879) in 100 mL of acetate buffer. The procedure in step IV was used to establish the standard curve using the following quantities: 0.00, 50, 250, and 500 nmol of glucose equivalents (Fig. 17b). V. Soluble sugars analysis using Anthrone Reagent  1.^Take the supernatants from step III and dilute them appropriately. As in the starch analysis, the dilution factor  118 will vary from one plant part to another, time of year, etc. For spruce try the following: Needles: 40 AL plant extract + 160 AL distilled water. Roots: 50 gL plant extract + 150 ML distilled water. Stems: 60 gL plant extract + 140 pL distilled water. 2.  To deproteinize, add to each of the above 50 mL each of barium hydroxide and zinc sulphate solutions (0.3 M).  3.  Centrifuge for 3 minutes, then pipet 200 AL of the supernatant into a test tube.  4.^To each of the above (step 3), add: - 200 AL of 11.6 M HC1 - 40 AL of 45% formic acid - 1.6 mL of 80% sulphuric acid with anthrone ^(i.e., 20 mg anthrone (Sigma A-1631) plus 100 mL of 80% sulphuric acid). For each day prepare fresh samples of 80% H 2 SO 4 and anthrone solution. 5.^Close the top of the test tube with a marble. Vortex very very gently and briefly (watch out for bubbles). 6.  Put the entire rack in a boiling water bath for exactly 12 minutes.  7.  After 12 minutes, remove the rack and put it in an ice water bath until cold (5 - 10 minutes).  8.  Vortex briefly. Let the samples sit in the ice water bath for 5 - 10 minutes to let the bubbles settle.  9.  Read the absorbance at 630 nmol. Refer to: JERMYN, M.A. 1975. Anal Biochem 68:332-335. YEMM, E.W. and WILLIS, A.J. 1954. Biochem 57: 508514. Glucose concentrations were calculated from a glucose standard curve. Prepare the glucose standard solution by dissolving 18 mg of D-glucose (BDH Inc. B10117) in 100 mL of distilled water. The procedure in step V was used to establish the standard curve using the following quantities: 0.00, 50, 100, 150, and 200 nmol glucose (Fig. 17a).  VI. Preparation of solutions used in the carbohydrate analyses  1.^Acetate buffer (150 mM) a) Weigh out 4.082 g of sodium acetate powder into a measuring cylinder. Add distilled water to 200 mL level and stir well.  ^  119 b) In a second cylinder, measure 2.58 mL of acetic acid. Then carefully add distilled water to 300 mL level and stir well. c) Mix a) and b) together. The pH of the mixture should be approximately 4.5. 2.^Amyloglucosidase a) Weigh out 800 units of amyloglucosidase powder (E.C.3.2.1.3, from Rhizopus, Sigma A-7255) and dissolve it in 1.0 mL of distilled water. b) Add 3-5 mg of charcoal, mix very well and then spin (desk top).^Pipet the supernatant and to it add 0.4228 g of granular ammonium sulphate. Mix gently. c) Spin pellet. and mix and save  (desk top). Pipet the supernatant out and keep the Re-suspend the pellet in 1.0 mL of distilled water gently. If you get some charcoal particles, re-spin the supernatant.  3.^a-Amylase a) Weigh out 40,000 units of a-amylase powder (E.C.3.2.1.1, from Aspergillus, Sigma A-0273) and dissolve it in 1.0 mL of distilled water. b) as in 2. b) c) as in 2. c) 4.^Peroxidase/glucose oxidase/colour reagent a) Enzymes Capsules containing 500 units of glucose oxidase (from Aspergillus niger), 100 Purpurogallin units of peroxidase (horseradish) and buffer salts can be purchased from Sigma (PGO Enzymes, cat. no. 510-6). Dissolve 1 capsule in 100 mL of distilled water in an amber bottle. Mix well by inverting the bottle several times. b) o - dianisidine dihydrochloride (colour reagent) Weigh out 50 mg of o-dianisidine dihydrochloride powder and dissolve it in 20 mL of distilled water. Mix well. c) To a) add 1.6 mL of the colour reagent solution from b) and mix well by inverting the bottle several times. For more details refer to Sigma Diagnostics Procedure No. 510. This leaflet accompanies kit 510-DA which comes with PGO Enzymes, colour reagent, glucose standard, and barium hydroxide and zinc sulphate solutions.  120 (A) GLUCOSE STANDARD CURVE.  S6^lee^150  ^  200  ^  250  GLUCOSE (nmol)  (B) STARCH STANDARD CURVE.  lie^atie^300^400  GLUCOSE EQUIVALENTS (nnol)  Figure 17. Glucose (a), and starch (b) standard "curves"  developed for carbohydrate analyses.  BIOGRAPHICAL INFORMATION  NAME: Bernard Malata Chomba  MAILING ADDRESS: Forest Department, Division of Forest Research  P.O. Box 22099, Kitwe. ZAMBIA.  PLACE AND DATE OF BIRTH: Mporokoso District, Zambia, Dec. 05, 1953.  EDUCATION (Colleges and Universities attended, dates, and degrees):  University of Dar-es-Salaam, Morogoro, Tanzania: B.Sc. (nor.) (Hons.) 1982 Zambia Forest College, Mwekera, Kitwe, Zambia: Dip. in Forestry, 1979  POSITIONS HELD:  Silviculturist/Fuelwood Project Manager: Jan. 1985 - Aug. 1989 Silviculturist: Jan. 1983 - Dec. 1984 Tree Improvement Forester: May - Dec. 1979  PUBLICATIONS (if necessary, use a second sheet):  (see attached sheet)  AWARDS:  1989 -1992: International Development Research Centre (IDRC, Ottawa, Canada) 1986: British Council Scholarship, Oxford, U.K. 1984: Finnish International Development Agency (FINNIDA, Helsinki, Finland) 1980 - 1982: World Bank/Zambia Government Scholarship 1977 - 1979: Zambian Government Forestry Diploma Scholarship Complete one biographical form for each copy of a thesis presented to the Special Collections Division, University Library.  PUBLICATIONS Chomba, B.M., and J. Saramaki. 1985. Coppice yield for short rotation Eucalyptus on the Copperbelt. Zambia Forest Department. Research Note 37. 16 p. Chomba, B.M., and A.C. Mubita. 1988. Three year results of a fuelwood research project in Zambia. Zambia Forest Department. Country Report. 52 p. Saramaki, J., and B.M. Chomba. 1985. Effects of pruning on the growth of Pinus kesiya. Zambia Forest Department. Research Note 35. 13 p. Selander, J., and B.M. Chomba. 1989. Control of Lantana camara L. in forest plantations. Zambia Forest Department. Research Note 44. 12 p. Selander, J., and B.M. Chomba. 1989. Eradication of stump regrowth in Eucalyptus grandis. Zambia Forest Department. Research Note 45. 11 p.  

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