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Proceedings of the National Academy of Sciences of the United States of America logoLink to Proceedings of the National Academy of Sciences of the United States of America
. 1999 Oct 26;96(22):12941–12946. doi: 10.1073/pnas.96.22.12941

Genetic ablation of root cap cells in Arabidopsis

Ryuji Tsugeki 1,*, Nina V Fedoroff 1,
PMCID: PMC23173  PMID: 10536027

Abstract

The root cap is increasingly appreciated as a complex and dynamic plant organ. Root caps sense and transmit environmental signals, synthesize and secrete small molecules and macromolecules, and in some species shed metabolically active cells. However, it is not known whether root caps are essential for normal shoot and root development. We report the identification of a root cap-specific promoter and describe its use to genetically ablate root caps by directing root cap-specific expression of a diphtheria toxin A-chain gene. Transgenic toxin-expressing plants are viable and have normal aerial parts but agravitropic roots, implying loss of root cap function. Several cell layers are missing from the transgenic root caps, and the remaining cells are abnormal. Although the radial organization of the roots is normal in toxin-expressing plants, the root tips have fewer cytoplasmically dense cells than do wild-type root tips, suggesting that root meristematic activity is lower in transgenic than in wild-type plants. The roots of transgenic plants have more lateral roots and these are, in turn, more highly branched than those of wild-type plants. Thus, root cap ablation alters root architecture both by inhibiting root meristematic activity and by stimulating lateral root initiation. These observations imply that the root caps contain essential components of the signaling system that determines root architecture.


The root cap is a structure that protects the root apical meristem, senses environmental signals, facilitates soil penetration by the root, and creates a chemical microenvironment (15). In Arabidopsis, the root cap develops from two sets of meristematic cells, a central group of initials that gives rise to the columella and a surrounding ring of cells that gives rise to both the lateral root cap and the epidermis (6). The columellar cells differentiate to produce amyloplasts, believed to be the statoliths whose gravitational sedimentation triggers root graviperception (7, 8). Peripheral root cap cells are secretory cells, producing and secreting mucilage as well as proteins and other metabolites (912). Root caps are highly dynamic structures, able to regrow and redifferentiate rapidly when removed or sloughed off as border cells (13, 14). Roots sense and respond to gravity by reorienting growth, and the root cap is the site of graviperception (1519). Roots also respond to other environmental stimuli, such as light, touch, and water, and there is evidence that the sensing mechanisms for some of these stimuli also reside in root caps (2024). Physiological changes in the root cap precede the gravitropic curvature of roots. Differential lateral transport of auxin and calcium have been observed in the root cap of horizontal roots before the onset of the differential growth that results in curvature (25). Recent molecular-genetic analysis of Arabidopsis mutants altered in gravitropism have led to the identification of both auxin influx and efflux carriers as well as other potential signal transduction molecules, supporting the hypothesis that auxin transport plays an important role in the root gravitropic response (2630).

Although the role of the root cap in the root gravitropic response is well documented, the developmental role of root caps has not been studied because it has not been possible previously to eliminate root caps throughout the plant’s life cycle. To determine whether the root cap plays a role in the development of the plant as a whole, we genetically ablated root cap cells by using a root cap-specific promoter to express a diphtheria toxin gene (31). The diphtheria toxin A-chain (DT-A) is known to kill cells by ribosylating the EF2 translation-initiation factor and inhibiting protein synthesis (32, 33). DT-A has been used to selectively ablate tissues and cells in higher plants, including Arabidopsis (3438). In the present work, we isolated an Arabidopsis root cap-specific promoter and used it to express the DT-AtsM gene. We obtained transgenic plants exhibiting the agravitropic phenotype expected for plants lacking root caps. Such plants have normal aerial parts but short, prematurely differentiated roots with disrupted root caps as well as more secondary and tertiary lateral roots than wild-type plants. These observations reveal that the root cap is an essential component of the signaling system responsible for maintaining root growth rates and suppressing lateral root formation.

Materials and Methods

Plant growth media were purchased from GIBCO/BRL, restriction enzymes from Boehringer Mannheim, and reagents for microscopy from Electron Microscopy Sciences (Fort Washington, PA). Common laboratory reagents were obtained from Sigma-Aldrich.

Plant Material.

Arabidopsis thaliana ecotype Nossen (No-0) was used in all experiments. The activator (Ac) (NaeAc 380–6) and dissociation (Ds) (Ds2 389–25) lines used here have been described (39, 40). Plants were grown on basic Murashige–Skoog (MS) agar medium containing 1% sucrose with or without one or more selective agents such as hygromycin and kanamycin. Growth conditions were 12 hours light, 12 hours dark, 22°C. Images of vertically grown seedlings or their roots were captured to a Macintosh computer by using a ProgRes 3012 charge-coupled device camera (Kontron, Zurich) and daguerre software and analyzed by using the NIH image analysis program (developed at the National Institutes of Health and available at http://rsb.info.nih.gov/nih-image/). The angle of root growth from the vertical gravity vector was determined by measuring the angle between the gravity vector and a line drawn between the root tip and a point about 2/3 of the distance between the hypocotyl and the tip.

Strains and Molecular Cloning.

Plasmids were grown by standard protocols in Escherichia coli strain DH5α (41). Agrobacterium tumefaciens strain ASE was used for Arabidopsis transformation. Total DNA was isolated from plants as described (42). Thermal asymmetric interlaced PCR (TAIL-PCR) was carried out as described, except that 2.5 units of AmpliTaq Gold (Perkin–Elmer, Roche Molecular System) was used in each reaction (42, 43). Genomic DNA fragments corresponding to the Ds insertion site were isolated from an No-0 genomic DNA library by using the 3′ TAIL-PCR fragment as a probe. A 1.4-kilobase (kb) ScaI fragment containing the putative promoter of the RCP1 gene was cloned into pBluescript KS+ at the HincII restriction site to generate pKSSS. The HindIII–SpeI fragment from pSR101 (constructed by S. Raina and N.V.F.) contains a β-glucuronidase (GUS) gene that has a nuclear localization signal followed by nopaline synthase (NOS) terminator (nlsGUS–NOS) and was cloned into pKSSS to generate pKSSGN. The Asp718–SpeI fragment from pKSSGN was then cloned into the pCGN1548 binary vector (Calgene, Davis, CA) to create pCGSGN (44).

The EcoRI–SpeI fragment containing the gene for the temperature-sensitive DT-AtsM from pSKDT-M, a gift of Hugo J. Bellen (Baylor College of Medicine, Houston), was filled in by using the Klenow fragment of E. coli DNA polymerase and cloned in pBluescript KS+ at the EcoRV restriction site to generate pKSDT. The NOS terminator EcoRI–SpeI fragment from pSR101 was cloned into pKSDT to generate pKSDN. The HindIII–SpeI fragment from pKSDN, which contains the DT-AtsM gene fused to the NOS terminator, was cloned into the pKSSS plasmid to generate pKSSDN. The Asp718–SpeI fragment from pKSSDN was then subcloned into the pCGN1548 binary vector, which was digested with Asp718 and XbaI, to create pCGSDN. The binary vector constructs, pCGSGN and pCGSDN, were used to transform Agrobacterium by the freeze-thaw method (45).

Plant Transformation.

Agrobacterium transformation of Arabidopsis with vacuum infiltration was carried out as described at http://www.bio.net/hypermail/ARABIDOPSIS/9606/0025.html. Kanamycin-resistant transformants were selected on MS agar medium containing 1% sucrose, 50 μg/ml kanamycin, 100 μg/ml carbenicillin, and 10 μg/ml benomyl.

Histochemical GUS Assays.

Seedlings and dissected roots were stained for GUS activity for 16–24 hours at 37°C in the 0.5 mg/ml X-Gluc (5-bromo-4-chloro-indolyl-β-d-glucuronide, Rose Scientific, Cincinnati)/100 mM sodium phosphate buffer, pH 7.0/10 mM EDTA/0.5 mM K4Fe(CN)6/0.5 mM K3Fe(CN)6/0.1% Triton X-100, and then fixed as described below.

Light Microscopy.

X-Gluc-stained tissues were prepared for whole mounts as described (46). For sectioning, GUS-stained seedlings or dissected roots were fixed for 4 hours at 4°C in 1% glutaraldehyde/4% paraformaldehyde in 50 mM sodium phosphate buffer (pH 7.0). Tissues were then washed in the same buffer, dehydrated in a graded ethanol series, infiltrated with Spurr’s resin, and polymerized. Unstained tissues were fixed and washed in the same way and then postfixed in 1% osmium tetroxide in 50 mM sodium phosphate buffer (pH 7.0) for 2 hours and dehydrated as described above, except for a final acetone wash, followed by infiltration with Spurr’s resin in acetone and subsequent polymerization. Sections (2–5 μm thick) were cut with the Ultra-Microtome MT-2 (Sorvall) equipped with a glass knife. The sections were dried onto glass slides. Samples not stained for GUS activity were stained with 1% toluidine blue-O in 1% sodium borate at 60°C for 30 seconds and then quickly rinsed with distilled water. Samples were visualized with Nomarski (differential interference contrast) optics on a Zeiss Axioskop microscope. Images were captured as described above and then processed by using Adobe photoshop software (Adobe Systems, Mountain View, CA) to create figures.

Results

Identification and Characterization of a Gene Expressed in the Root Cap.

The root cap-specific promoter used in the present experiments was initially identified in an enhancer-trap line (102-1) obtained by using the previously described Ac/Ds transposon tagging system (39, 40, 42, 47). The Ds transposon carries a promoterless GUS reporter gene that can be activated by insertion into or near a promoter or enhancer. GUS gene expression in roots of line 102-1 commences several hours after germination and increases during early development (Fig. 1 AD). GUS activity is detectable only in root cap cells, and the gene is expressed in the root cap cells of lateral and adventitious roots, as well as those of the primary root.

Figure 1.

Figure 1

(AD) GUS-stained primary root tips of line 102-1 at 0 hour (imbibed) (A), 10 hours (B), 1 day (C), and 4 days (D) after transfer to 22°C. (E) Longitudinal section of a stained primary root tip of a transgenic plant expressing the GUS gene from the root cap-specific promoter. (F) The columella and lateral root cap cells expressing the GUS gene in E are indicated by large and small red circles, respectively. (Bar = 20 μm.)

The sequences flanking the Ds insertion site were amplified from total DNA of line 102-1 by TAIL-PCR (42, 43). Several fragments were amplified and sequenced, including a 1.2-kb, a 1.0-kb, and an 0.6-kb fragment extending from the 5′ end of the Ds, as well as a 1.2-kb fragment extending from the 3′ end of the Ds. All of the amplified fragments contained sequences identical to those either at the 5′ or the 3′ end of the Ds followed by genomic DNA sequences. Each Ds end was flanked by an identical 8-bp sequence. Because insertion of Ac/Ds transposons causes a duplication of 8 bp at the insertion site, it follows that the TAIL-PCR fragments are derived from the sequences flanking the 5′ and 3′ end of a single Ds insertion (48). This was confirmed by genomic DNA blot hybridization, which showed that line 102-1 contains a single Ds-homologous insertion and that it hybridizes to both the 5′ or 3′ TAIL-PCR fragments (data not shown).

The 3′ flanking sequence was almost identical to that of an Arabidopsis ecotype Columbia cDNA expressed sequence tag (1,439 bp in length; GenBank accession no. AF168390). The expressed sequence tag was used to clone the gene from an Arabidopsis (ecotype Nossen) genomic library. The gene, designated RCP1 for root cap 1 (4,438 bp in length; GenBank accession no. AF168391), comprises nine exons and eight introns. It encodes an ORF of 415 aa with no significant similarity to any gene in the GenBank database. The Ds is inserted just upstream from the putative initiator ATG of the ORF, and there is a stop codon in frame 15 bp upstream of the initiation codon (Fig. 2A). Based on genomic DNA blots probed with gene-specific probes, the RCP1 gene is unique in the Arabidopsis genome (data not shown).

Figure 2.

Figure 2

The Ds insertion site and a diagram of the constructs used for Arabidopsis transformation. (A) Sequence at the Ds insertion site. The putative initiator ATG of the RCP1 gene is shown in boldface. The 8-bp Ds target site duplication is underlined by arrows. The underlined TAA is a stop codon in-frame with the initiator ATG. The 5′ end of the RCP1 expressed sequence tag cDNA is indicated by a bent arrow at the stop codon. One of the ScaI restriction sites used to isolate the root cap promoter is shown. (B) The 1.4-kb ScaI promoter fragment of the RCP1 gene was fused to a GUS gene containing nuclear localization signal (nlsGUS) and the DT-AtsM gene, each of which is followed by a transcriptional terminator (NOS) taken from a gene for nopaline synthase. ScaI* indicates the location of the restriction site shown in A.

To determine whether the expression of RCP1 gene is abolished as a result of the Ds insertion, an RNA gel blot containing poly(A)+ RNA isolated from roots and leaves of wild-type and Ds-insertion line 102-1 plants was probed with the RCP1 expressed sequence tag cDNA. The probe hybridized to two RNA species whose sizes were about 1.4 kb and 0.9 kb in RNA samples prepared from both roots and leaves of wild-type and Ds-insertion line 102-1 plants (data not shown). Detection of hybridizing RNAs in Ds-insertion line 102-1 raised the possibility that the expression of the RCP1 gene is not entirely abolished by insertion of the Ds transposon. This possibility is strengthened by the recent observation that the 3′ end of the Ds element has weak promoter activity (K. Scortecci, R. Raina, N.V.F., unpublished data). In addition, because the insertion of the Ds is not in a coding region of the RCP1 gene, expressed RNA may be able to produce a functional protein.

Root Cap Promoter.

Although the Ds-disrupted RCP1 gene is expressed in both roots and leaves, the enhancer-trap 102-1 line shows GUS expression only in the root cap cells. Thus, the genomic sequence upstream of the Ds insertion site appears not to include elements required for expression in leaves. To determine whether the sequence upstream of the Ds insertion site contains a root cap-specific promoter, we tested its ability to direct expression of a reporter gene to root cap cells. The 1.4-kb ScaI fragment ending 23-bp upstream from the Ds insertion site was fused to a GUS gene containing a nuclear localization signal (Fig. 2B) and the construct was introduced into Arabidopsis. Transgenic lines expressed the GUS gene only in root cap cells, both of primary roots (Fig. 1E) and of lateral roots. Although some staining is detectable in the layer of initials (Fig. 1 E and F), it is not uniform across the cells, suggesting that it is attributable to diffusion. Hence, the 1.4-kb upstream sequence contains the root cap-specific promoter.

Genetic Ablation of Root Cap Cells.

We carried out genetic ablation of root cap cells to examine the developmental consequences of their absence. The 1.4-kb upstream sequence that confers expression of the GUS gene in the root cap cells, subsequently referred to as the root cap promoter, was used to drive expression of DT-AtsM (31). Because DT-AtsM is a cell-autonomous toxin, only cells expressing DT-AtsM are expected to be killed. The root cap promoter was fused to the DT-AtsM gene followed by the NOS terminator (Fig. 2B). This construct was introduced into Arabidopsis plants by using Agrobacterium-mediated transformation. We obtained 22 independent kanamycin-resistant transgenic lines, of which 13 lines had short, agravitropic roots. The rest of the transgenic lines were indistinguishable from untransformed plants. Three of the 13 lines with a short-root phenotype and four lines of the transformed but phenotypically normal lines were crossed with the original Ds-insertion line to assess the effect of toxin gene expression on the GUS expression pattern. All of the F1 plants from crosses to short-root plants also had short roots, and no GUS staining was detected in the tips of either primary or lateral roots. All of the F1 progeny from crosses with transgenic but phenotypically normal plants had normal roots and GUS staining in root caps (data not shown). Thus, a majority of the transgenic plants containing the toxin gene exhibited a dominant, genetically transmissible phenotype that includes the production of short, agravitropic roots in which root cap-specific expression of the GUS gene is abolished. We conclude that expression of the DT-AtsM gene from the root cap promoter is responsible for these phenotypic effects and that expression of the toxin gene abolishes root cap-specific gene expression and root cap function.

Although temperature sensitivity of the DT-AtsM protein has been reported in other eukaryotic systems, including plants, we were not able to detect temperature sensitivity of the short-root or agravitropic phenotypes (31, 38, 49). The presence of the DT-AtsM gene is correlated with the short-root, agravitropic phenotype irrespective of growth temperature in the 16–28°C range. We conclude that the enzyme is expressed but that its activity is either not temperature-sensitive in root cap cells of Arabidopsis in the temperature range tested or that enough activity persists at the nonpermissive temperature to eliminate root cap function.

Histological Analysis of the Root Tips of DT-AtsM Transgenic Plants.

We carried out a comparative histological analysis of the root tips of 1-, 2-, and 4-day-old DT-AtsM transgenic and wild-type plants to assess the effect of toxin gene expression on root-tip structure. The radial organization of roots in the transgenic lines was unaltered (Fig. 3). However, two layers of columella root cap cells were entirely missing, and the remaining three layers of cells were increasingly disorganized as the root developed. In addition, some of the lateral root cap cells were missing. The cells in the root tips of transgenic plants were bigger than those in wild-type roots, and there were fewer cytoplasmically dense cells, suggesting that the short-root phenotype of DT-AtsM plants is attributable to a reduction in root meristematic activity. The cells of the quiescent center, which are usually mitotically inactive and remain small during root growth, were also larger in transgenic DT-AtsM plants than in wild-type plants. Despite the overall decrease in mitotic activity, division of quiescent center cells was occasionally observed (Fig. 3D). Development of epidermal root hairs, vascularization, and vacuolization commence much closer to the initials in DT-AtsM than in wild-type roots (Fig. 4). Vacuolization is known to commence earlier in non-root-hair cells than in root-hair cells of Arabidopsis (6). Such differential vacuolization in root-hair and non-root-hair cells also appears to occur in the transgenic plants but in closer proximity to the root tip than in wild-type roots (Fig. 4). These observations show that expression of the DT-AtsM gene alters root differentiation in addition to disrupting root cap structure and function.

Figure 3.

Figure 3

Longitudinal sections of primary root tips. One-day-old root tips of wild-type (A) and transgenic (D) plants; 2-day-old root tips of wild-type (B) and transgenic (E) plants; 4-day-old root tips of wild-type (C) and transgenic (F) plants. Division of quiescent cells was occasionally observed in transgenic roots, as seen in D (arrow). (Bar = 20 μm.)

Figure 4.

Figure 4

Differentiation of root cells in transgenic and wild-type plants. Wild-type (A and B) and DT-AtsM (C and D) seedlings were grown on MS agar medium for 4 days. (Bar = 50μm.)

Root Growth Is Altered in Transgenic DT-AtsM Plants.

We have compared the root growth patterns of transgenic DT-AtsM and wild-type plants. The primary roots of the transgenic plants grew very slowly, attaining lengths of only 4–6 mm in 11 days compared with a length of 40–50 mm for wild-type roots under the same conditions (Fig. 5). Moreover, the roots of DT-AtsM plants were completely agravitropic. The direction of primary root growth is plotted in Fig. 6 for wild-type and DT-AtsM transgenic seedlings. Unlike wild-type roots, the roots of transgenic plants show no preference for growth along the gravity vector, indicating that they are totally lacking the gravitropic response.

Figure 5.

Figure 5

Root growth rates. Seedlings were grown on vertical MS agar plates. The mean values of roots of wild-type (No-0) and DT-AtsM plants were plotted (n = 32 and 32 roots, respectively).

Figure 6.

Figure 6

Direction of primary root growth. Wild-type and DT-AtsM (SSD) seedlings were grown on the surface of a vertical MS agar plate for 7 days. The angle of the primary root from the vertical was determined and plotted as a point on the circular graph. The numbers inside the graph are the numbers of plants whose root angles from the vertical are within each 30° range.

The distribution of the number of lateral roots on primary roots and secondary (lateral roots on lateral roots) and tertiary lateral roots (lateral roots on secondary lateral roots) is shown in Fig. 7 for 14 day-old wild-type and transgenic plants. The average numbers of secondary lateral roots in the transgenic lines tested were 10.2 and 13.2 for lines 2-1 and 6-5, respectively, compared with 1.9 for wild-type plants. Moreover, only 1 of 69 (1.4%) of the wild-type seedlings had tertiary lateral roots, whereas 28 of 61 (46%) and 25 of 55 (45%) DT-AtsM transgenic seedlings of lines 2-1 and 6-5, respectively, produced tertiary lateral roots. As a result, although the number of lateral roots in the transgenic plants is lower than that in wild type, they are more highly branched, having more secondary and tertiary lateral roots. Hence, continuous root cap ablation accelerates formation of lateral roots, implying that the root cap is either the source of a signal that inhibits or a sink for a signal that stimulates lateral-root development.

Figure 7.

Figure 7

Distribution of the number of lateral roots. Wild-type and DT-AtsM seedlings were grown on MS agar medium for 14 days. The number of lateral roots on 69 wild-type plants (No-0) and on 61 (line 2-1) and 55 (line 6-5) transgenic plants was counted under a dissecting microscope. Each histogram shows the distribution of the number of primary lateral roots, secondary lateral roots, and tertiary lateral roots. The sum of the number of primary, secondary, and tertiary lateral roots was for each wild-type and DT-AtsM transgenic plant (total).

The Aerial Parts of Transgenic DT-AtsM Plants Are Normal.

Despite the abnormal root structure of DT-AtsM transgenic lines, the appearance of the aerial parts of the transgenic plants was normal both on MS agar medium and in soil. This may be because the formation of more lateral roots compensates for the effect of the short-root phenotype.

Hormone Treatment Does Not Rescue the Short-Root Phenotype.

The effect of plant hormones on root meristematic activity was tested by adding 2,4-dichlorophenoxyacetic acid, indole-3-acetic acid, 6-benzylaminopurine, gibberellic acid, and 1-aminocyclopropane-1-carboxylic acid (a precursor of ethylene) to the medium on which plants were grown. None of the growth regulators was able to restore the root growth rates of transgenic plants. Thus, development of the root system in DT-AtsM transgenic lines could not be normalized by the exogenous supply of the growth regulators tested, indicating that the defect is not simply a quantitative one.

Discussion

In previous experiments, root caps have been eliminated by surgical removal or laser ablation, establishing that root caps are essential for root graviperception (50). Because root caps regenerate rapidly, the developmental consequences of root cap elimination cannot be assessed in such experiments. To assess their developmental role, we genetically ablated Arabidopsis root caps by expressing a DT-A gene from a root cap-specific promoter. Transgenic plants expressing the DT-AtsM gene produced morphologically normal shoots and highly abnormal roots. As expected, roots of the transgenic plants were agravitropic. But in addition, the roots grew much more slowly, differentiated prematurely, and showed extensive secondary and tertiary branching. Cytological analyses revealed that the cellular organization of the root tip was not extensively altered, although the root caps had fewer cell layers and the remaining cells were disorganized. Observed changes in cell size and cytoplasmic density imply that the elimination of root caps reduces mitotic activity in the root tip.

It has been reported previously that laser ablation of Arabidopsis and surgical removal of maize root cap cells has no effect on root elongation rate (19, 50). In other studies, it was observed that removal of maize root caps transiently stimulates root growth (51, 52). Genetic ablation of root caps in the present study inhibited primary root growth strongly. However, genetic ablation suppresses root cap production continuously, whereas root cap function is rapidly restored after cell ablation or physical removal of the root cap. Thus, when the central cells in the root apical meristem were laser-ablated, their fragmented remains were displaced toward the root tip by proximal vascular cells, which rapidly ceased expressing a vascular cell-specific marker and began to express a root cap-specific marker instead, indicating that the cells had changed fate in response to their new position within the root (13). In the present experiments, any cell that acquires the capacity to express the root cap-specific promoter will be killed by expression of the DT-AtsM gene. Continuous root cap ablation also accelerates lateral-root formation. It was previously known that removal of the root tip, including the root apical meristem and the root cap, stimulates formation of lateral roots (53). The results of the present experiments demonstrate that elimination of just the root caps suffices to stimulate lateral-root formation. Although we do not know whether the cumulative effect of root cap loss on root meristematic activity is primary or secondary, it appears most likely that genetic ablation of the root caps disrupts cell signaling.

Both root elongation and lateral-root formation require auxin transport, and there is evidence for both acropetal and basipetal movement of auxin within the root (5458). Lateral-root formation depends on auxin. It is stimulated by either supplying auxin exogenously or by introducing genes that increase the internal auxin concentration (53, 59). Conversely, lateral-root formation is inhibited in auxin-insensitive mutants (6062). The source of the auxin that promotes lateral-root formation is the shoot. Radiolabeled auxin applied to shoots accumulates in lateral-root primordia, and blocking auxin transport from shoots to roots inhibits lateral-root formation (58, 63). Although it is known that formation of lateral roots can be influenced by environmental conditions, these differences may be mediated by auxin redistribution as well (6466). Thus, the most straightforward hypothesis to explain the observed changes in root architecture in the transgenic DT-AtsM plants is that essential components of the auxin redistribution system reside in the root cap. A reduction in the root growth rate consequent on destruction of the transport system by genetic ablation is consistent with the observation that root growth can be inhibited by auxin-transport inhibitors (56, 57). Stimulation of lateral-root formation by root cap ablation suggests that auxin continues to move from shoots to roots in the transgenic plants and accumulates in the capless roots. This, in turn, suggests that there is an auxin sink in the root cap or that access to the sink is through the root cap auxin-transport system. Finally, the observation that plants lacking root caps produce normal aerial parts suggests that the putative root cap auxin transport/sink system is not coupled with the shoot’s auxin-transport system or the root’s acropetal auxin-transport system.

Acknowledgments

We thank Dr. Hugo J. Bellen (Baylor College of Medicine) for providing us a gene for the temperature-sensitive DT-A, Dr. Surabhi Raina and Dr. Ramesh Raina (Pennsylvania State University) for stimulating discussion, Rosemary Walsh and Michelle Peiffer (Pennsylvania State University) for valuable suggestions for sectioning, and Xi-Ying Yu and Maya L. Olson (Pennsylvania State University) for assisting in growing plants. Sectioning was performed at the Electron Microscopy Facility at the Pennsylvania State University. This work was supported by a grant (NAG5-3970) from the National Aeronautics and Space Administration (NASA) to N.V.F.

Abbreviations

Ac

Activator

Ds

Dissociation

DT-A

diphtheria toxin A-chain

MS

Murashige–Skoog

TAIL

thermal asymmetric interlaced

NOS

nopaline synthase

GUS

β-glucuronidase

kb

kilobase

Footnotes

Data deposition: The sequence reported in this paper has been deposited in the GenBank database (accession no. AF168391).

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