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Physiology and Development |
Department of Environmental, Population, and Organismic Biology, University of Colorado, Boulder, Colorado 80309-0334 USA
Received for publication May 14, 2002. Accepted for publication April 17, 2003.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Key Words: Acomastylis rossii • carbon allocation • labeled carbon pulse chase experiment • preformation • Rosaceae • resource allocation • southern Rockies, USA
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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; Bloom et al., 1985
). Patterns of growth determine, in part, patterns of carbon allocation, yet current ideas about carbon allocation for many plants are based upon incomplete knowledge of development. For example, current understanding of carbon dynamics and development in herbaceous perennials of the alpine tundra is based largely on observations of aboveground phenology. Most tundra perennials are leafless throughout the winter; they expand and mature leaves within a few weeks of snowmelt, after which no additional leaves mature until the following year. Leaves function for a single growing season and then senesce in the autumn. It has been assumed that the initiation and development of preformed buds, containing primordia that will mature the following year, occurs at the end of a growing season after assimilation has largely been completed (e.g., Mooney and Billings, 1960
; Mark, 1970
; Jaeger and Monson, 1992
). Preformed organs then mature the following spring. These phenological observations and assumptions have led to the conclusion that there is a pronounced asynchrony between carbon supply and demand in tundra perennials (Bliss, 1960
; Mooney and Billings, 1960
; Fonda and Bliss, 1966
; Billings, 1987
; Chapin et al., 1990
).
Recent studies of alpine perennials, however, have revealed that existing concepts of alpine plant development, in which primordia are initiated in the fall as aerial organs are senescing, are not broadly applicable to all tundra plants (Aydelotte and Diggle, 1997
; Diggle, 1997
; Meloche and Diggle, 2001
). Initiation and growth of preformed leaf primordia have been shown to begin as soon as snow melts in the spring and to proceed throughout the entire growing season. Consequently both initiation and growth of primordia occur simultaneously with photosynthesis in mature leaves. Development of organ primordia also can continue long after the aerial organs have senesced. Furthermore, growth of leaf and inflorescence primordia within the apical bud can take from one to four full years from initiation to maturation and function, depending upon the species (Sørensen, 1941
; Mark, 1970
; Aydelotte and Diggle, 1997
; Diggle, 1997
; Meloche and Diggle, 2001
).
Given that initiation, morphogenesis, and growth of organ primordia occur throughout the photosynthetic season and beyond, new analyses that integrate carbon allocation and developmental dynamics are required. The species Acomastylis rossii (Rosaceae) is an ideal candidate for examination of carbon allocation to preforming organs in an herbaceous tundra perennial. Acomastylis rossii is abundant and widespread in the alpine tundra of the southern Rockies (Walker et al., 1993
). The development of this species has been studied in detail (Meloche and Diggle, 2001
); Acomastylis rossii undergoes extensive preformation with each leaf and inflorescence developing over a period of 3 yr (Fig. 1). In addition, the leaves of this species reach up to one-third of final mature length as primordia still within the apical bud and can contain as much as 20% of mature leaf biomass. Thus, carbon allocated during preformation is a significant fraction of mature leaf biomass.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). Acomastylis rossii is a dominant component of the vegetation in both moist and dry meadow communities in the southern Rockies.
Plants grow monopodially and produce a branched underground rhizome system, making it difficult to identify distinct individuals (Spomer and Salisbury, 1968
). Therefore, for the purposes of this study, an "individual" is defined as the rosette of leaves generated by a single shoot apical meristem, inflorescences subtended by those leaves, and 4–6 cm of unbranched rhizome subjacent to the apical meristem. The individual rhizomes always bear roots at or near the point where the shoot emerges from the ground, and each inflorescence is subtended by a functioning leaf (C. G. Meloche, personal observations). Based on these observations, it is inferred that each "individual" is an integrated physiological unit sensu Watson and Casper (1984)
and capable of affecting its own uptake, storage, growth, and sexual reproduction. Also supporting this conclusion are data from preliminary labeling experiments that showed newly applied carbon is not transported more than 6 cm back from the shoot apical meristem (C. G. Meloche, unpublished data). Typically rhizomes grow only 3 mm/yr. Thus, a 4–6 cm rhizome segment encompasses tissue up to 20 yr old.
Acomastylis rossii has been the subject of several studies of phenology and physiology (Mooney and Billings, 1960
; Holway and Ward, 1965
; Spomer and Salisbury, 1968
; Walker et al., 1994
). Acomastylis rossii accumulates stored starch in rhizomes, and the relative abundance of various carbohydrate fractions in different portions of the plant body changes throughout the growing season (Mooney and Billings, 1960
). The morphological development of this species also has been studied in detail (Meloche and Diggle, 2001
; Fig. 1). Acomastylis rossii undergoes extensive preformation with each leaf and axillary inflorescence developing over a period of 3 yr. Organs are initiated and develop belowground in the apical bud for 2 yr then emerge aboveground and function in the third year. Defining a cohort as all of the organs initiated in a single growing season, each individual bears one cohort of mature leaves and one cohort of leaf primordia that had been initiated the previous season (cohorts A and B in Fig. 1). In addition, leaf initiation begins at or prior to snowmelt and a third cohort of leaf primordia are added throughout the growing season (cohort C in Fig. 1). Prior to dormancy, at the end of a growing season, there is one cohort of senesced leaves aboveground and two cohorts of leaf primordia belowground in the apical bud. Previously, plants have been assumed to enter dormancy when the aboveground leaves have lost all green coloration, between mid-August and mid-September (Mooney and Billings, 1960
; Holway and Ward, 1965
). However, we have shown that development continues belowground within the apical bud until late October (Meloche and Diggle, 2001
). For the purposes of this study leaf senescence is considered separately from plant dormancy. Leaves were considered senesced when they lacked any green coloration upon visual inspection.
Study site
Labeling and harvesting took place in the alpine tundra of Niwot Ridge (elevation 3750 m) in the Front Range of the Colorado Rocky Mountains (40°03' N, 105°36' W). Detailed climatic records have been maintained for this site since 1981 (Losleben, 1983–1988
). Experiments were performed on individuals growing in a fellfield, characterized as being dominated by Silene acaulis (L.) and Trifolium dasyphyllum (T. & G.), and in a moist meadow codominated by A. rossii and Deschampsia caespitosa (L.) (May and Webber, 1982
).
13CO2 labeling
In order to determine the source of carbon for preforming organs (see Fig. 2), 32 haphazardly selected plants in each of the two communities were exposed to an atmosphere enriched in 13CO2 (modified from Chiariello et al. [1989]
). At the same time, 32 control plants were selected at random within the same areas. At the time of labeling all plants had one cohort of mature leaves that had been fully expanded for 3–5 wk. Each experimental individual was enclosed in a large sealed plastic bag. A cup containing 0.5 g of pure 13C sodium carbonate (Sigma-Aldrich, St. Louis, Missouri, USA) and a Pasture pipette containing an excess of acetic acid were sealed into each bag. Before dawn the acid was added to the sodium carbonate, releasing 13CO2 into the bag. Plants were exposed to the 13CO2-enriched atmosphere for one photoperiod on 13 July 1998. The bags were removed after sunset on the same day. Conditions on that day were clear with maximum, minimum, and mean temperatures of 17.0°, 9.0°, and 12.6°C, respectively. Total solar radiation for the day was 30.12 x 106 J/m2. Winds were from the north-northwest at a mean speed of 4.63 m/s (Losleben, Niwot Ridge LTER data; http://culter.colorado.edu:1030/Niwot/NiwotRidgeData/Saddle.html).
After labeling, plants were harvested at intervals over the ensuing 13 mo in order to trace the movement of the label. Harvests of labeled plants and unlabeled controls took place on the following dates: 3 wk after treatment on 3 August 1998; after the end of the photosynthetic season on 25 September 1998; at the beginning of the second growing season on 25 June 1999; and after the end of the second growing season on 6 September 1999. Sample size was eight plants per site per date for both label and control treatments. After each harvest, plants were dissected into the following constituent morphological compartments: previous years senescent leaves, mature leaves, leaf primordia, rhizome, and roots. The rhizome was further divided into the youngest 1 cm of rhizome (which included all attachment points of leaf primordia, the current year's mature leaves, as well as the subjacent 3 yr of growth), and the subjacent 1 cm of rhizome. This division of the rhizome is intended to test for attenuation of the label at increasing distance from the mature leaves. In addition, for harvests performed in 1999 the primordia were divided into those that were initiated in 1998 and those initiated in 1999. This subdivision is necessary to distinguish between primordia present when the label was applied and those initiated after labeling, and hence, to discriminate among the models (Fig. 2B vs. 2C). Tissue samples were oven dried at 80°C for 48 h.
Mass spectrometry
Dried samples were ground to a particle size of <250 µm using a stainless steel ball mill (Wig-L-Bug, Crescent Dental, Chicago, Illinois, USA). Two-milligram subsamples were packaged in 5 x 8 mm tin capsules and carbon isotopic composition was analyzed by means of isotope ratio mass spectrometry. Mass spectrometer analysis was performed on a Europa Scientific Intergra IRMS (PDZ Europa, Crew, Cheshire, UK) at the University of California, Davis, Stable Isotope Facility, Davis, California, USA.
Statistical analyses
The natural abundance of 13C in each morphological compartment was calculated from control plants collected from the study sites at each time interval and was then subtracted from the levels present in labeled plants. This net level of 13C enrichment above background levels was used in all subsequent analyses. Thus, reported 13C values are specifically quantities of label, not total 13C. Eight treated individuals were significantly damaged during the course of the experiment and were therefore excluded from further analyses. Values of 13C enrichment from labeled plants growing in the moist meadow and the fellfield were pooled after relative abundance of the label in the morphological compartments in the two communities was determined to be indistinguishable (data not shown). Descriptive statistical analyses of 13C enrichment and t tests were performed using Statview 4.5 for the Macintosh (Abacus Concepts, 1993
). Multiple means were compared by ANOVA using the GLM procedure in JMP 4 (Academic Version; SAS Institute). Data are primarily reported as total mass distribution of the label to indicate patterns of allocation (Boutton, 1991
). Concentration in micrograms of 13C per gram of dry mass is included in some instances to indicate the sink strength of particular morphological compartments.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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The mean total quantity of 13C label recovered from plants harvested in June of 1999 was 427.0 ± 108.8 µg (Fig. 3C). This label was most abundant in the senesced leaves that had been mature in 1998; they contained 187.0 ± 49.5 µg (40.6%) of recovered 13C. Mature, functioning leaves contained 114.3 µg ± 28.6 (28.4%) of the label. Inflorescences that were mature in 1998 contained only 9.3 ± 1.6 µg (2.2%) of the label. Inflorescence stalks of A. rossii tend to shatter in the winter, and most of the biomass contained in this compartment is unrecoverable under field conditions. There were no mature inflorescences produced by any of the labeled individuals in June 1999. The stems and roots averaged 82.8 ± 18.7 µg (24.5%) and 25.3 ± 8.8 µg (5.4%) of label, respectively. The leaf primordia remaining in the apical bud in June retained 8.0 ± 1.4 µg (2.7%) of the label. The leaf primordia present at this time constitute a very small biomass, hence the small percentage of the label. However, if the 13C levels are expressed as micrograms of label per gram of tissue, then the concentrations of label in primordia and mature 1999 leaves are similar. Mature leaves average 495.7 ± 99.4 µg of label per gram of tissue and primordia contain 391.3 ± 74.9 µg of label per gram of tissue. The difference between these values is not statistically significant (t = 0.839; df = 26, P = 0.409).
At the final harvest, after senescence of leaves in September 1999, the average amount of label recovered per plant was 469.0 ± 87.7 µg (Fig. 3D). The senesced leaves that had been mature and green at the time of labeling (1998 leaves) still retained the most label (Fig. 3D). The current year's (1999) mature leaves, also senesced by this time, had 135.4 ± 34.7 µg (28.9%) of the label. For this harvest, the two cohorts of leaf primordia present in the apical bud were analyzed separately. The older cohort of primordia had been present in 1998, at the time of labeling, whereas the younger cohort was initiated in 1999 and was never directly exposed to a 13C-enriched environment. The 1998 cohort averaged 1.1 µg (0.2%) of the added 13C and the leaf primordia initiated in 1999 contain 0.1 µg (0.01%) of the label. The quantity of label in these youngest leaf primordia is very small as is the concentration of the label per gram of dry mass. Of the total mass of carbon in the youngest cohort only 7 µg/g can be attributed to the label. By contrast, 80 µg/g of the carbon in the youngest segment of stem can be attributed to the label, an order of magnitude higher concentration.
In order to fully summarize the pattern of variation in 13C label, the data were subjected to ANOVA with time (sampling date) and morphological compartment as main effects (Table 1). Although the mean total quantity of 13C declined over each sampling interval, the main effect of time was not significant. In contrast, the mean quantity of 13C differed significantly among compartments. The significant interaction term indicates that the pattern of change in 13C over time differed among compartments (Table 1; Fig. 5). Variation over time in the individual compartments was analyzed further by separate one-way ANOVA (Table 2). Mean total quantity of 13C in roots and stems does not change over time, whereas variation in 13C levels of 1998 leaves and 1999 primordia over time is significant (Table 2; Fig. 5).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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; Diggle, 1997
; Meloche and Diggle, 2001
) indicates that current concepts of the timing of carbon allocation in alpine plants should be reconsidered. The observed pattern of 13C distribution among organs over two years demonstrates that preforming leaf primordia of A. rossii do use carbon from concurrent photosynthesis in mature leaves for growth and morphogenesis. Substantial quantities of 13C were detectable in developing leaf primordia in the first harvest after label was applied (Fig. 3A), indicating that carbohydrate is typically translocated directly to preforming organs without passing through a long-term storage pool. Furthermore, the high concentrations of 13C in the primordia demonstrate that these developing organs are a strong sink for newly fixed carbon (Fig. 4). Direct allocation of the label to the preforming organs eliminates the maximum asynchrony scenario (Fig. 2A). Preformation does not depend solely upon stored reserves.
To determine whether preforming organs of A. rossii use stored reserves of carbohydrate as well as concurrently fixed photosynthate for growth, the distribution of the label in the year following the pulse is critical (Fig. 2B vs. 2C). Is label that initially had been allocated to storage during year 1 reallocated to the development of newly initiated organ primordia in year 2? This analysis is complicated by the fact that leaf primordia spend 2 yr developing within the apical bud (Fig. 1). Plants harvested in August of the second year of the experiment contained both leaf primordia that had been present at the time of labeling (1998) and leaf primordia that were initiated in the second year. The older primordia (initiated in 1998) acquired label at the time of application, as described above, but the new primordia (initiated in 1999) could only acquire label from reserves stored in other parts of the plant. Primordia initiated in the second year did contain a small but significant quantity of label above the background concentration of 13C (one-sample t test, H0: µ = 0, P = 0.03, df = 7).
The presence of a small quantity of label in new primordia initiated during the second year suggests a minor role for carbohydrate that had been fixed and stored mid-season in the initiation and growth of preformed leaf primordia. Moreover, if storage in the starch-rich roots and rhizomes was the primary source of carbon for primordial development, then the concentration of label in the new primordia should be similar to that in these putative storage organs. However, the concentration of label found in the primordia was 10 times less than that in the roots and rhizomes. The very small 13C signature found in the newest primordia is most likely attributable to dilution of a small quantity of carbon supplied by storage organs in larger quantities of unlabeled carbon coming from concurrent photosynthesis.
An alternative explanation is that the presence of 13C in the youngest leaf primordia is due to contamination. The two cohorts of primordia are not delineated from one another by any morphological marker. They were separated for analysis based on the average number of leaves per cohort. Therefore, it is possible that some primordia from the older cohort were inadvertently included with the newest cohort and that there was actually no translocation of carbon from storage. Although we cannot distinguish between these scenarios, we did not detect a quantity of label in the newest primordia consistent with a large contribution of stored carbon from the roots and rhizomes. Furthermore, the roots, young rhizome segments, and older rhizome segments all failed to showed a significant decrease over time in the average total quantity of label that they contained (Table 2; Fig. 5). Together these results indicate that development of preformed organ primordia of A. rossii depends primarily on current photosynthate at the time when photosynthetic leaf area is at its maximum and does not typically utilize large quantities of carbon stored in the roots or rhizome when plants are actively fixing carbon.
Development using concurrently fixed carbon can account for much of primordium development when mature leaves are present. However, there are still substantial periods of time in the early spring and again in the late fall when growth takes place without the benefit of mature, photosynthetic leaves as a direct carbon source. This growth must utilize carbon input from storage. As a cohort of leaves matured in the spring of the second year of the study, there was a large increase in their total quantity of 13C compared to that detected in the same cohort of leaf primordia the previous fall (compare primordia in Fig. 3B with 1999 leaves in Fig. 3C; Fig. 5 1998 primorida). Unexpectedly, this increase in label was not accompanied by a significant decrease in the total quantities of the label in the putative storage organs, the roots and stems (Table 2; Fig. 5). If the 13C did not come from storage in the rhizome, what is the source of this carbon?
The large variance in the 13C data may have precluded detection of real changes in belowground carbon storage. Thus, additional study of the role of these organs in supporting primordium growth during early spring and late fall is needed. We suggest, however, that the preceding year's apparently senesced leaves should also be considered as a possible source of carbon moving into primordia late in the growing season. Because leaves of A. rossii do not abscise at senescence, we could measure, in the second year, the quantity of label in senesced leaves that had been mature at the time of labeling. The detected loss of label from senesced leaves from September 1998 to June 1999 averaged 225 ± 72 µg (1998 leaves in Fig. 5). Over the same time interval the mean total loss of label from an entire plant (164 ± 72 µg) was not significant (Table 1). We infer that after the September 1998 sampling date, carbon was translocated away from leaves that were photosynthetic in 1998 and, upon casual observation, appeared to have senesced. The lack of change in the amount of label in the roots and rhizomes suggests that carbon from the "senesced" leaves was not allocated to long-term storage in these structures. In contrast, the average quantity of 13C in developing leaf primordia increased over the September to June time interval. It is possible that the 13C recovered from senesced leaves was allocated, either directly or indirectly, to the growth of leaf primordia. An a posteriori contrast of the change in 13C level of senesced 1998 leaves with developing 1999 primordia over the September to June time interval was significant (F1,12 = 15.68, P < 0.001, see also Fig. 5). 13C levels in the two cohorts of leaves are changing in opposite directions, a pattern that provides support for the possibility that carbon can be reallocated from apparently senesced leaves. Developmental analysis has shown that initiation and growth of primordia continues throughout September and October in A. rossii (Meloche and Diggle, 2001
). These processes are potentially immediate sinks for carbon recovered from the senescing leaves; that is, carbon could be translocated directly from the senescing 1998 leaves to the 1999 leaf primordia in the late fall. Alternatively, carbon from senescing leaves could be allocated directly to storage in roots and rhizomes and then remobilized into expanding leaves in the spring, prior to the June sampling.
Extensive translocation of carbon out of leaves at or just before the time of senescence has been documented in numerous plants including deciduous trees (Gibbs, 1940
; Mooney and Hays, 1973
; McLaughlin and McConathy, 1979
; McLaughlin et al., 1980
; Abrusrewil et al., 1983
; Keller and Loescher, 1986
, 1989
; Smith et al., 1986
; Nguyen et al., 1990
; Kozlowski, 1992
), monocarpic herbs (Heilmeier et al., 1986
; Tissue and Nobel, 1990
; Tworkoski, 1992
; Minoletti and Boerner, 1993
), and herbaceous perennials (Mooney and Billings, 1960
; Miller and Rose, 1992
; Wyka, 1999
). We have found no reports that indicate how long translocation continues relative to the condition of leaves. Further study is required to determine how long apparently senesced leaves remain physiologically capable of transport.
Patterns of radiocarbon allocation have been examined in a few herbaceous perennials: Carex bigelowii, Podophyllum peltatum (Jónsdóttir and Watson, 1997
), and Lathyrus sylvestris (Magda et al., 1993
). These studies show that, despite differences in relative amounts of carbon allocated to storage vs. new shoot growth, the development of shoot primordia depends upon concurrently fixed carbon. The herbaceous perennials examined are phylogenetically and ecologically diverse, yet show similar direct allocation of carbon to developing shoot primordia. Acomastylis rossii, with far more extensive preformation, shows a similar relationship between development and carbon allocation. More taxa must be studied in order to understand whether general or predictable relationships between plant morphology, life form, and the patterns and timing of carbon allocation can be recognized.
The ubiquity of preformation among alpine plants has been variously attributed to time constraints imposed by a short growing season, poor growing conditions of the alpine environment, and a low-risk growth strategy of these plants (Sørensen, 1941
; Billings and Bliss, 1959
; Bliss, 1962
; Billings and Mooney, 1968
; Mark, 1970
; Diggle, 1997
). Such explanations interpret this developmental pattern as simply a negative consequence imposed by a harsh environment. By studying carbon allocation within a developmental context, we show that in Acomastylis rossii, preformation is associated with synchronization of carbon supply and carbon demand. By synchronizing carbon supply and demand, resources can be allocated directly to developing organs, thus reducing the cost of building dedicated storage structures and repeatedly translocating resources to and from a storage location.
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FOOTNOTES |
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2 Author for reprint requests (diggle{at}spot.colorado.edu
; Tel: 303-492-4860)
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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