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First published online July 28, 2006; 10.1104/pp.106.082594 Plant Physiology 142:207-219 (2006) © 2006 American Society of Plant Biologists OPEN ACCESS ARTICLE
A Standardized Method for Analysis of Medicago truncatula Phenotypic Development1,[W],[OA]United States Department of Agriculture, Agricultural Research Service, Department of Agronomy and Plant Genetics, University of Minnesota, St. Paul, Minnesota 55108 (B.B., C.P.V.); United States Department of Agriculture, Agricultural Research Service, Midwest Area Biometrician, Peoria, Illinois 61604 (D.P.); and Australian Research Council Centre of Excellence for Integrative Legume Research, Australian Research Council Centre for Complex Systems, Advanced Computational Modelling Centre, University of Queensland, Brisbane, Queensland 4072, Australia (J.H.)
Medicago truncatula has become a model system to study legume biology. It is imperative that detailed growth characteristics of the most commonly used cultivar, line A17 cv Jemalong, be documented. Such analysis creates a basis to analyze phenotypic alterations due to genetic lesions or environmental stress and is essential to characterize gene function and its relationship to morphological development. We have documented morphological development of M. truncatula to characterize its temporal developmental growth pattern; developed a numerical nomenclature coding system that identifies stages in morphological development; tested the coding system to identify phenotypic differences under phosphorus (P) and nitrogen (N) deprivation; and created visual models using the L-system formalism. The numerical nomenclature coding system, based on a series of defined growth units, represents incremental steps in morphological development. Included is a decimal component dividing growth units into nine substages. A measurement component helps distinguish alterations that may be missed by the coding system. Growth under N and P deprivation produced morphological alterations that were distinguishable using the coding system and its measurement component. N and P deprivation resulted in delayed leaf development and expansion, delayed axillary shoot emergence and elongation, decreased leaf and shoot size, and altered root growth. Timing and frequency of flower emergence in P-deprived plants was affected. This numerical coding system may be used as a standardized method to analyze phenotypic variation in M. truncatula due to nutrient stress, genetic lesions, or other factors and should allow valid growth comparisons across geographically distant laboratories.
Medicago truncatula has become a model plant for the study of legume biology (Cook, 1999  ). With the anticipated sequencing of its genome by 2007, its large expressed sequence tag database, and availability of the Medicago Affymetrix GeneChip, essential resources are available to genetically dissect this organism. However, to fully utilize genomic approaches, the Medicago research community requires standardized methods for growth and phenotypic analysis that can be utilized worldwide. A standardized system, similar to that adopted by the Arabidopsis (Arabidopsis thaliana) research community, should allow researchers from geographically distant regions to easily and clearly monitor morphological and developmental changes resulting from mutations, biotic and abiotic stress, and other factors, thereby facilitating the standardization of experimental results across laboratories.
The establishment of a standardized baseline for the temporal developmental growth pattern of M. truncatula should provide a uniform approach to compare growth of genetic mutants or ecotypes having a developmentally impaired phenotype. The development of a numerical nomenclature coding system provides a means to define specific growth stages in plant development, thereby making it possible to discern and document specifically where changes occur for phenotypically distinct plants. Additionally, a nomenclature coding system may be used to clearly communicate developmental stages without extensive descriptions. It also provides a basis to standardize tissue collection for analysis. A coding system may also be incorporated into a component of the M. truncatula plant ontology database, specifically the plant morphological aspect of the database (Blake, 2004
We have used nutrient stress to simulate changes in whole plant development to assess the effectiveness of a standardized coding system to reproducibly detect morphological changes in plant growth. We have followed shoot and root development of M. truncatula from cotyledon to early pod formation when grown under nitrogen (N)- and phosphorus (P)-deficient conditions. These particular stress conditions were chosen because of their major role in plant growth and their implication in nutrient signaling. P, an important plant macronutrient, is immobile in the soil and is a limiting nutrient in many agricultural systems (for review, see Raghothama, 1999 The overall goal of this work was to develop a standardized method to characterize the morphological development of M. truncatula, line A17 of cv Jemalong. Specifically, our objectives were: (1) to evaluate the temporal developmental vegetative growth pattern of M. truncatula under defined conditions; (2) to develop a standardized nomenclature coding system that identifies specific stages in morphological development; (3) to create a developmental structural model of M. truncatula that allows phenological and geometric data, from different experiments, to be visualized for ease of comparison; and (4) to validate the use of the nomenclature coding system to identify phenotypic differences when plants are grown under P- and N-stress conditions. This work was meant to create a framework for the future analysis of phenotypic alterations of M. truncatula due to either genetic mutations or environmental conditions.
The Numerical Nomenclature Coding System M. truncatula produces a procumbent, trailing type of growth habit from the combined elongation of the main and axillary shoots. As shoot apices develop, they produce a series of growth units defined as metamers that consist of an internode, leaf, and axillary bud. The main shoot, the primary axis of vegetative growth, initially produces 4 metamers separated by very short immeasurable internodes. Afterward, the internodes of the successively developing metamers elongate, producing additional trifoliate leaves with measurable internodal distances. A spiral type of phyllotaxy is produced in the elongating portion of the shoot. The axillary bud of each metamer has the potential to grow into an axillary shoot. The initiation and growth of these axillary shoots is coordinated with the elongation of the main shoot.
Based on the coding system developed for the quantitative modeling of Arabidopsis by Mundermann et al. (2005) The nomenclature coding system starts with the metamer associated with the unifoliate leaf as metamer 1 (m1). The units developing above this are numbered in ascending order with m2 associated with the first trifoliate, etc. The metamers forming axillary shoots developing off of the main axis are coded first according to their main shoot metamer of origin then numbered sequentially starting at the base with 1 (e.g. the axillary shoot associated with m1 that contains 1 unit of growth is designated as m1-1). Floral location is indicated with an "F" preceding the number of its associated metamer of emergence. Flowers emerging from the axils of the main shoot are designated according to the metamer of origin followed by F0 (e.g. floral emergence from m6 is designated as m6-F0). When considering a flower developing along an axillary shoot, it is coded according to the main shoot metamer of origin followed by the axillary shoot metamer of emergence preceded by "F" (e.g. the flower emerging from the axillary shoot associated with m1 and from the axil of m3 of the m1 axillary shoot is coded as m1-F3). The numerical coding system identifying units of M. truncatula growth and their location is illustrated in Figure 1 . Additionally, the nomenclature coding system defines leaf and reproductive organ development into nine substages (Table I ) depicted by a decimal added as an extension to the numerical code. These substages were chosen because they are visually identifiable and provide a continuum of growth and development for the organ that they specify. Therefore, to indicate that the m1-1 leaf is fully developed, a code of m1-1.9 is used. A measurement component is also associated with the coding system. Various parameters were measured that could distinguish growth alterations that may be missed by the coding system alone. Plant growth parameters measured for this study included shoot and root fresh and dry weights, shoot and root total lengths, leaf size, and internode lengths. There are numerous other parameters that may have been measured throughout development. The above parameters were chosen because they are nondestructive and augment the morphological focus of this study.
By following plant growth under nutrient stress conditions, specifically N and P stress, we have simulated potential changes in whole plant development that may occur due to genetic lesions or other factors. This allowed us to demonstrate the use of the coding system and its measurement component to document plant morphological development. Moreover, by using these treatments, we were able to assess the effectiveness of this system to distinguish differences in phenotypic alterations. When shoot tissue of P-deficient and control plants were analyzed for percentage of P, plants grown under P-deficient conditions had a lower P content relative to the controls (compare 0.194% P, SE ± 0.014, n = 3 for P-deficient plants, and 0.462% P, SE ± 0.042, n = 3 for control plants). The low P tissue content in P-deprived plants indicates that the morphological alterations reported in this study can be attributed to the P stress imposed. The results from this study were also used to construct an empirical model of M. truncatula growth and development under these stress conditions. Animations showing model development can be viewed online in Supplemental Figures 1 and 2.
A simple way to orient oneself to the developing plant is to use the unifoliate leaf as a reference marker. The unifoliate leaf, associated with m1, emerges from the developing apical meristem between the two cotyledons (Fig. 2A ). The second leaf (associated with m2) emerges 180° opposite to the unifoliate. The developing leaves from m1 and m2, plus the cotyledons, produce a cross type of appearance to the developing plant (Fig. 2, A–C). The third leaf (associated with m3) develops 140° relative to the m2 leaf, adjacent to the m1 leaf. The fourth leaf (associated with m4) develops on the same side of the cotyledons as the m2 leaf and approximately 165° from the m3 leaf (Fig. 2D).
Figure 2 shows metamer location and orientation at key developmental stages of plant growth for the three nutrient treatments (Fig. 2, A–H). Figure 2 also shows the model rendition of some of the key developmental stages plus an empirical reconstruction of plants at 40 d after planting (dap; Fig. 2, I–K). Figure 3 depicts a time line for the chronological progression of plant growth throughout the stages of development over a 40-d period. It includes the time interval to reach key developmental stages and the correlation of these stages with the onset of axillary shoot development and floral emergence. Table II presents a summary of shoot temporal development and the numerical code associated with growth for the three nutrient treatments. It compares growth stage codes and the time it takes to reach these stages. It also shows statistical comparisons of the decimal code for the nutrient-deficient plants relative to plants grown under nutrient-sufficient conditions. An animation of shoot temporal development showing the sequential appearance of shoot structures in a color-coded manner can be viewed online in Supplemental Figure 2.
By 7 dap, there were no visible differences in plant development between the three nutrient treatments (fig. not shown). All plants had fully expanded cotyledons (this occurs by 4 dap) that had a succulent characteristic and were blue-green in color. m1 started to develop as evidenced by the presence of the unifoliate leaf at a mean developmental stage of m1.4, indicating that the unifoliate leaf blade is folded and the petiole is visible (Table I).
By 14 dap, plants grown under both nutrient-sufficient and P-deficient conditions had m1 at full development or near full development (m1.9 and m1.8, respectively) and m2 at a mean developmental stage of m2.7 (Fig. 2, A and B; Table II). However, N-deficient plants began to show a significant delay in development with m1 and m2 (Fig. 2C; Table II; F-test, P
By 21 dap, the leaf of m3 control plants had a mean developmental stage of m3.9 (Fig. 2D; Table II). Under P-deficient conditions, this leaf was at a mean developmental stage of m3.7 (Fig. 2E; Table II). This is the first sign of a significant developmental difference between nutrient-sufficient and P-deficient plants as detected by the nomenclature coding system (F-test, P By 28 dap, the leaf associated with m5 was at a mean developmental stage of m5.9 for plants grown under nutrient-sufficient conditions and at m5.7 for P-deficient plants (Fig. 2, G and H; Table II). It should be noted that by 28 dap, the cotyledons of P-deficient plants were clearly chlorotic and some showed signs of senescence. For N-deficient plants, m5 was not visible until 32 dap (Fig. 3; Table II). By 32 dap, the leaf associated with m6 for control plants was at a mean developmental stage of m6.9, and for P-deficient plants it was at a developmental stage of m6.7. The main shoot developed subsequent metamers between 32 and 40 dap. By 40 dap, a total of 10 to 11 metamers were produced along the main shoot of both nutrient-sufficient and P-deprived plants. In contrast, the N-deficient plants produced only five metamers (Fig. 3; Supplemental Fig. 1).
The first axillary bud to break and initiate growth was associated with m1 (Fig. 3; Table II). The m1 axillary bud break was initiated between 18 and 21 dap in plants grown under nutrient-sufficient conditions (Fig. 3; Fig. 2D) and was correlated with the development of m4 along the main shoot. The m1 axillary bud break of P-deficient plants occurred between 21 and 25 dap and was correlated with the development of m5 along the main shoot (Fig. 3). By 40 dap, the axillary shoot of m1 was a mean developmental stage of m1-6.4 and m1-5.5 for plants grown under nutrient-sufficient and P-deficient conditions, respectively. N-deficient plants failed to develop any axillary shoots. It should be noted that axillary shoot elongation began to occur when the third leaf emerged from its growing point. The second axillary bud to break and initiate growth was associated with m2 (Fig. 3; Table II). The m2 axillary bud break was initiated between 21 and 25 dap in plants grown under nutrient-sufficient conditions and was correlated with the development of m5 along the main shoot. The m2 axillary bud break of P-deficient plants occurred between 25 and 28 dap and was correlated with the development of m6 along the main shoot (Fig. 3). By 40 dap, the axillary shoot of m2 had a mean developmental stage of m2-6.3 and m2-5.4 for plants grown under nutrient-sufficient and P-deficient conditions, respectively. The third axillary bud to break and initiate growth was associated with m3. The m3 axillary bud break was initiated between 25 and 28 dap for plants under nutrient-sufficient conditions and was correlated with the development of m6 along the main shoot (Fig. 3; Table II). The m3 axillary bud break of P-deficient plants occurred between 28 and 32 dap and was correlated with the development of m7 along the main shoot (Fig. 3). By 40 dap, the m3 axillary shoot was at a mean developmental stage of m3-5.4 and m3-4.4 for nutrient-sufficient and P-starved plants, respectively. Subsequently, the m4 axillary shoot developed between 28 and 32 dap for plants under nutrient-sufficient conditions and between 32 and 40 dap for P-deficient plants (Fig. 3; Table II). Concurrently, m8 and m9 developed along the main shoot. By 40 dap, the m4 axillary shoot was at a developmental stage of m4-4.4 and m4-3.4 for nutrient-sufficient and P-deficient plants, respectively. Axillary bud break from m5 was initiated by 32 dap in control plants and between 32 and 40 dap in P-deficient plants (Fig. 3; Table II). Axillary bud break from m5 was correlated with the development of m9 and m10 along the main shoot. By 40 dap, the m5 axillary shoot was at the m5-3.4 and m5-2.3 developmental stage for nutrient-sufficient and P-deficient plants, respectively. Axillary buds from m6 and m7 initiated growth just prior to 40 dap for both nutrient conditions (data not shown). By 40 dap, the m6 and m7 axillary shoots contained 1 leaf.
Flowers began to emerge along the main shoot by 32 dap in both nutrient-sufficient and P-deficient plants (Fig. 3; Table II). However, P-deficient plants had approximately 3-fold fewer flowers that initially emerged at 32 dap, and these flowers were at a significantly younger developmental stage relative to plants grown under nutrient-sufficient conditions (compare mean flower developmental stage of 0.1–0.3, respectively; n = 11; t test, P < 0.05; Fig. 4A ; Table II). Notably, by 40 dap, no differences in number of emerging flowers or their developmental stage was evident for both treatments (Fig. 4A; Table II). The differences in flower number between the nutrient-sufficient and P-deficient plants may be attributed to the delayed development of P-stressed plants because the timing of flower emergence and not position is affected between the two treatments. The initial location of flower emergence for both nutrient-sufficient and P-deficient plants was from the axils of m5, m6, and m7. As the main shoot continued to elongate, flowers emerged from axils above, but not below, these metamers. N-deprived plants failed to produce any visible reproductive structures.
On elongated axillary shoots, flowers were visible at 32 dap on control plants and at 40 dap on P-deficient plants (Table II). Flowers emerged from elongated axillary shoots that originated from m1, m2, m3, and m4 for both nutrient-sufficient and P-deficient plants. However, plants grown under nutrient-sufficient conditions had nearly 2-fold more flowers that developed from the elongated axillary shoots relative to P-stressed plants (Fig. 4B). Similar to flower emergence along the main shoot, differences in the number of emerged flowers on the elongating axillary shoots may be attributed to the delayed developmental stage of axillary shoots in P-stressed plants relative to the controls. The location of flower emergence along the axillary shoots occurred at a specific metamer from the main shoot metamer of origin and was different for the m1 axillary relative to the other axillary shoots. The first flower to have emerged for the m1 axillary was from the m1-3 position, whereas the first flower to have emerged from the other axillary shoots is from the m2-2, m3-2, and m4-2 positions (Table II). Differences in flower developmental stage between control and P-deficient plants was evident for flowers that emerged from the axillary metamer closest to the main shoot where flowers from the control plants were at a slightly advanced developmental stage relative to the P-deficient plants (Table II). However, such a delay in flower emergence may also be attributed to a delay in axillary shoot development for P-deficient plants. Subsequently, it was difficult to assess whether the delay in flower emergence along the main and axillary shoots for P-deficient plants could be attributed solely to a delay in whole plant development or if a physiological response to P stress could also contribute to such a response.
Growth parameters for total shoot length, internode length of main and axillary shoots, shoot and root fresh weights, and leaf dimensions were measured every 4 to 7 d beginning at 7 dap. These measurements were used to augment the nomenclature coding system, making the system more sensitive to the identification of phenotypic differences associated with variable treatments.
Main shoot total length and internode lengths were measured at 40 dap for plants grown under nutrient-sufficient and P-deficient conditions. The N-deficient plants failed to produce measurable shoot length and therefore were not included in this analysis. The results showed no significant differences for total length of the main shoot for plants grown under nutrient-sufficient and P-deficient conditions (t test, P
Measurement of axillary shoot total length at 40 dap showed that axillary shoots were significantly longer in plants grown under nutrient-sufficient relative to P-deficient conditions (t test, P
These results indicate that P deficiency had a greater affect on axillary shoot elongation relative to main shoot elongation. Such a response resulted in P-deficient plants appearing stunted relative to plants grown under nutrient-sufficient conditions. The effect of P deficiency on axillary shoot elongation was not evident until 28 dap and does not affect internodes produced before this time. Therefore, prior to 28 dap, M. truncatula appeared to contain sufficient P to sustain growth comparable to control plants. Whether the lack of difference in main shoot total length would be significant at later stages remains to be established. Perhaps the main shoot was more tolerant of P stress relative to the axillary shoots.
Leaf petiole length, blade length (the length of the middle leaflet including its petiolule), and blade width (distance across the two lateral leaflets including their petiolules) were measured for fully developed leaves associated with m1 through m5 along the main shoot (Fig. 6, A and B ).
Leaf petiole lengths were significantly different between the three treatments (F-test, P  0.05; Fig. 6A). Differences in petiole length between plants grown under nutrient-sufficient and N-deficient conditions were evident at the m1.9 stage of development. By comparison, differences in petiole length of nutrient-sufficient and P-deficient plants were evident at the m2.9 stage. Petiole length progressively increased as leaves were produced along the main shoot for nutrient-sufficient and P-deficient plants. In contrast, petiole length for N-deficient plants did not vary and remained short relative to the other treatments (Fig. 6A).
Significant differences in blade length were also detected between the three nutrient treatments (F-test, P Differences in blade width were not detected between nutrient-sufficient and P-deficient plants (data not shown). For N-deficient plants, blade width was difficult to assess because leaf blades failed to completely open.
Root architectural development of M. truncatula grown under nutrient-sufficient, P-deficient, and N-deficient conditions was analyzed over a 28-d period. Plants were harvested at 4, 7, 14, 21, and 28 dap. Roots were excised and scanned on a flatbed scanner to evaluate root architectural development. Measurements for total length of primary root, first-order laterals, second-, and higher-order laterals were calculated. Additionally, total root number for first-order, second-, and higher-order laterals was also determined. Table III shows a numerical summary of root growth over a 28-d period as compared to shoot growth stages. Figure 7 illustrates this data in a graphical format. Digital scans comparing M. truncatula root architectural development for plants grown under nutrient-sufficient, P-deficient, and N-deficient conditions over a 28-d period can be viewed online in Supplemental Figure 3.
By 7 dap, short first-order lateral roots were visible on all plants regardless of nutrient treatment. There were no differences between the three nutrient treatments when considering primary root length and first-order lateral root total length and total root number (Fig. 7, A–C; Table III).
By 14 dap, second-order lateral roots developed on all plants regardless of nutrient treatment. Second-order lateral root total length and total root number was significantly lower in the N-deficient plants relative to the other nutrient treatments at this time (F-test, P
By 21 dap, no differences were detected in first-, second-, and higher-order lateral root total length and number between nutrient-sufficient and P-deficient plants (Fig. 7; Table III). However, for N-deficient plants, first-order lateral root length was significantly shorter and total root number was significantly lower relative to the other two nutrient treatments (F-test, P
By 28 dap, differences between nutrient-sufficient and P-deficient plants became visible (Fig. 7, D and E; Table III). Plants grown under P-deficient conditions showed root total length and number lower for second- and higher-order laterals relative to plants grown under nutrient-sufficient conditions (F-test, P Shoot-root ratios, based on tissue dry weights, were evaluated at the completion of the study. Shoot-root ratios were similar for plants grown under nutrient-sufficient and P-deficient conditions with a value of 0.9. However, the shoot-root ratio of plants grown under N-deficient conditions was lower with a value of 0.55. Root hair characteristics were compared between the three nutrient treatments (Fig. 8 ). Differences in root hair appearance, relative to control plants, were first visible at 14 dap for N-deficient and at 21 dap for P-deficient plants. Root hairs of plants grown under P-deficient conditions (Fig. 8B) were longer relative to their nutrient-sufficient counterparts (Fig. 8A). Additionally, in P-deficient plants, root hair development occurred closer to the root tip in many instances. Root hairs of N-deficient plants were similar in appearance to those of control plants (Fig. 8C). However, the frequency of appearance along the roots varied between the two treatments. Roots of N-deficient plants had a glabrous appearance, with root hairs either lacking or patchy in distribution.
The M. truncatula growth analysis and model presented here provide a standardized method to evaluate phenotypic development. In this report, we have: (1) defined a detailed, reproducible baseline description of the temporal growth and developmental pattern of M. truncatula from cotyledon to early pod formation; (2) developed a standardized numerical nomenclature coding system that uses easily identifiable developmental growth stages to define plant growth; (3) identified plant morphological differences resulting from growth under nutrient stress conditions, thereby illustrating how the nomenclature coding system can be used to discern phenotypic alterations; and (4) developed an empirical model of M. truncatula development that captures differences in plant structure due to the different nutrient treatments for visualization purposes and provides a framework for future modeling of the genetic and/or physiological mechanisms underlying M. truncatula growth and development.
The detailed description of vegetative growth and reproductive organ emergence makes it possible to monitor morphological growth and development because it evaluates plant structures that persist and are visible to the unaided eye. Previous characterizations of M. truncatula focused on floral and pod traits (Lesins and Lesins, 1979
We have created a numerical nomenclature coding system, based on the system outlined by Mundermann et al. (2005) We have validated the effect of the coding system and its measurement component in detecting morphological changes in growth due explicitly to treatment effects over morphological plasticity. Using nutrient stress to modify whole plant development, we simulated changes that may occur due to genetic lesions or other factors. Growth under N and P deprivation produced a number of morphological alterations that could clearly be documented using both the coding system and measurement component. Specifically, N- and P-deprived plants showed a delay in leaf development and expansion along the main and axillary axes of growth (Figs. 2, 3, and 6; Table II), a delay in axillary shoot emergence and elongation (Figs. 2, 3, and 5; Table II), a decrease in leaf and shoot size, and an alteration in root growth (Figs. 5, 6, and 7; Table III). Furthermore, the timing and frequency of flower emergence in P-deprived plants were also affected (Fig. 4; Table II). Both nutrient stress conditions produced similar growth effects; however, the timing, extent, and location of these alterations differed depending upon the nutrient stress imposed.
Most of our results were compatible with and could be integrated into previously published studies that addressed the physiological, biochemical, and genetic mechanisms of plant responses to nutrient stress. When the description of morphological development is integrated with the mechanistic data and viewed in a comprehensive manner, the relationship between function and morphological development can be better realized. For example, the sequential timing of shoot, root, and reproductive structure development of M. truncatula grown under P deprivation was similar with reports on P and carbon cycling and remobilization in a variety of plant systems, including Glycine max, Stylosanthes hamata, Phaseolus vulgaris, Triticum aestivum, and Oryza sativa (Fredeen et al., 1989
However, not all of our results were consistent with the existing literature. For example, root architectural analysis of M. truncatula under P deficiency deviated from that reported for Arabidopsis or P. vulgaris, where low P conditions initially increased lateral root length (Lynch and Brown, 2001
Timing of development in this study has been described in terms of dap, with measurements taken at 4- to 7-d intervals. This means that our models cannot reflect continuous growth of plant parts in detail, as is possible if more frequent measurements are made (Mundermann et al., 2005
An aspect of timing requiring further study is the potential for incorporating the effects of temperature as a driver of physiological processes (Johnson and Thornley, 1985 In conclusion, our results demonstrate the application of a reliable numerical nomenclature coding system plus its measurement component to reveal altered phenotype and development resulting from growth under N and P deficiency. We have demonstrated that our standardized nomenclature coding system describing morphological alterations can be used in conjunction with other results, making it useful for comparative analyses across laboratories and thereby leading to a better understanding of the relationship between plant function and whole plant morphological development. Taken together, the integration of various techniques to study whole plant development can only lead to a more comprehensive understanding of the mechanisms driving the growth of M. truncatula.
Medicago truncatula seeds of line A17 of cv Jemalong were chemically scarified with concentrated sulfuric acid for 8 min and surface sterilized for 3 min with commercial-grade bleach (5.25% sodium hypochlorite). Seeds were given a 2-d germination period at 4°C then transferred to petri plates with moistened filter paper and given a 14-d vernalization period at 4°C. Vernalized seeds with a radical length of 1 to 1.5 cm were planted in pots containing quartz sand. Pot size was 10 cm x 10 cm x 35 cm. Plants were grown at 24°C to 26°C and a 16-h photoperiod (http://www.isv.cnrs-gif.fr/embo01/manuels/pdf/module1.pdf) with a light intensity at the sand surface of 430 to 490 µmol m–2 s–1. Plants were fertilized 3 times a week (150 mL of nutrient solution per fertilizer application) with the following nutrient solutions: complete nutrients, final concentration: KNO3, 15 mM; Ca(NO3)2 4H2O, 12.5 mM; Ca(H2PO4)2, 1 mM; MgSO4 7H2O, 1 mM; Fe EDTA, 0.01 mM; MnCl2, 0.004 mM; H3BO3, 0.02 mM; ZnSO4 7H2O, 0.0004 mM; NaMoO4, 0.0001 mM; and CaSO4 5H2O, 0.0001 mM. Nutrient solution lacking P substituted Ca (H2PO4)2 with 0.74 mM CaSO4 2H20. Nutrient solution lacking N substituted KNO3 and Ca (NO3)2 4H2O with 12.5 mM K2SO4 and 9 mM CaSO4 2H2O. Data were collected at 4, 7, 11, 14, 18, 21, 25, 28, 32, and 40 dap. Five seedlings for each treatment were selected for continuous monitoring of shoot growth. These five plants were photographed and various parameters were measured throughout the 40-d period. Plant growth parameters measured included shoot and root fresh and dry weights, shoot and root total lengths, leaf size, and internode lengths. An additional set of plants was grown in parallel for collection and processing of tissue. Total P analysis was done on dried shoot tissue of P-deficient and control plants harvested at 40 dap. Tissue P content was analyzed by optical emission spectroscopy. Three replicated experiments were performed. Images were taken with a Sony DSC-D770 digital camera. Growth angles were calculated from the digital images. All other parameters were measured using a handheld ruler.
An empirical model of M. truncatula development capturing differences in plant structure due to the different nutrient treatments was developed using the L-system-based L-studio software from the University of Calgary (Prusinkiewicz et al., 2000
To statistically examine developmental scale differences between the treatments, the Levene's test was first performed to satisfy the parametric ANOVA test assumption of homogeneity of variance and normality. A single-factor ANOVA was used to analyze developmental scale treatment differences. If a significant F-test statistic was obtained, then comparisons between each treatment and the control were performed using a Dunnett's test. Additionally, t tests for unequal replication and equal or unequal variances were used for the analysis of leaf and flower developmental scale differences and shoot and internode length differences. For most analyses, differences were considered to be significant at P
We thank Becky Schirmer for her technical assistance, Sue Miller for her helpful suggestions, and Michael P. Russelle for his valuable discussions concerning this study. Received April 26, 2006; accepted July 12, 2006.
1 This work was supported by the U.S. Department of Agriculture, Agricultural Research Service Current Research Information System (CRIS no. 3640–21000–019–00D), and by the Australian Research Council.  The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Carroll P. Vance (vance004{at}umn.edu).
[W] The online version of this article contains Web-only data.
[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.106.082594 * Corresponding author; e-mail vance004{at}umn.edu; fax 651–649–5058.
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