Generation of Signaling Speciﬁcity in Arabidopsis by Spatially Restricted Buffering ... · Generation of Signaling Speciﬁcity in Arabidopsis by Spatially Restricted Buffering

Generation of Signaling Specificity in Arabidopsis by SpatiallyRestricted Buffering of Ligand–Receptor Interactions C W OA

Emily B. Abrash,1 Kelli A. Davies,1 and Dominique C. Bergmann2

Department of Biology, Stanford University, Stanford, California 94305-5020

Core signaling pathways function in multiple programs during multicellular development. The mechanisms that compart-

mentalize pathway function or confer process specificity, however, remain largely unknown. In Arabidopsis thaliana,

ERECTA (ER) family receptors have major roles in many growth and cell fate decisions. The ER family acts with receptor

TOO MANY MOUTHS (TMM) and several ligands of the EPIDERMAL PATTERNING FACTOR LIKE (EPFL) family, which play

distinct yet overlapping roles in patterning of epidermal stomata. Here, our examination of EPFL genes EPFL6/CHALLAH

(CHAL), EPFL5/CHALLAH-LIKE1, and EPFL4/CHALLAH-LIKE2 (CLL2) reveals that this family may mediate additional ER-

dependent processes. chal cll2 mutants display growth phenotypes characteristic of er mutants, and genetic interactions

are consistent with CHAL family molecules acting as ER family ligands. We propose that different classes of EPFL genes

regulate different aspects of ER family function and introduce a TMM-based discriminatory mechanism that permits

simultaneous, yet compartmentalized and distinct, function of the ER family receptors in growth and epidermal patterning.

INTRODUCTION

The orchestration of programs that produce, pattern, and protect

developing tissues is a complex regulatory problem.Not onlymust

individual cells execute complex genetic programs to obtain dis-

tinctive identities and morphologies, but they must do so in con-

cert with their neighbors. Both plants and animals use peptide

ligands and cell surface receptor-based signaling to establish cell

fate and pattern and to regulate self-renewal. In plants, this

signaling is often mediated by leucine-rich repeat, single trans-

membrane pass receptor-like kinases (LRR-RLKs; Shiu and

Bleecker, 2001). Putative peptide ligands for these LRR-RLKs

are structurally diverse, ranging in size from modified pentapep-

tides to small (50 to 100 amino acids long) proteins and are often

part of large gene families (reviewed in Butenko et al., 2009; Jun

et al., 2010; Rowe and Bergmann, 2010; Rychel et al., 2010). The

roles and receptor interactions of these putative ligands remain

largely unknown, as do the mechanisms that enable related

ligands to function in distinct processes and interact with different

receptors.

One of the most extensively studied LRR-RLKs is that encoded

by theArabidopsis thalianaERECTA (ER) gene,mutations inwhich

result in altered plant stature and morphology (Torii et al., 1996).

Demonstration of ER functions in development (cell proliferation,

cell fate specification, and tissue patterning inmost aerial organs),

in homeostasis, and in immunity indicate thatER is a central player

in multiple signaling contexts (Godiard et al., 2003; Xu et al., 2003;

Shpak et al., 2004, 2005; Masle et al., 2005; Pillitteri et al., 2007).

ER is partially redundant with two close paralogs, ERECTA-LIKE1

(ERL1) and ERL2, and these three genes are known as the ER

family (ERf). Although ERL1 and ERL2 are capable of substituting

for ER when expressed in its place, their endogenous expression

patterns are distinct from that of ER, and they confer clear loss-of-

function stomatal and growth phenotypes only in an er back-

ground (Shpak et al., 2004, 2005).

The patterning of stomata (epidermal pores that facilitate

plant/atmosphere gas exchange) provides a simple, sensitive,

and quantifiable system for the study of basic developmental

phenomena. InArabidopsis, asymmetric, stem cell–like divisions

and cell–cell signaling are required for the arrangement of

stomata in stereotyped patterns, and stomata and precursors

can be analyzed to provide qualitative and quantitativemeasures

of signaling outputs (Bergmann and Sack, 2007). For example,

elimination of the entire ER family (er erl1 erl2) results in dramatic

overproduction of mispatterned stomata (Shpak et al., 2005).

Examination of lineage progression defects in single and double

mutants suggested that ER is primarily responsible for restricting

entry divisions, whereas ERL1 and ERL2 maintain the prolifera-

tive capacity of intermediate precursor stages (Shpak et al.,

2005). Stomatal production and pattern are also regulated by a

LRR receptor-like protein (LRR-RLP), TOO MANY MOUTHS

(TMM) (Nadeau and Sack, 2002). Elimination of TMM leads to

stomatal overproliferation and mispatterning phenotypes in leaves

similar to, but less severe than, those of er erl1 erl2 (Geisler et al.,

2000; Nadeau and Sack, 2002). Curiously, loss of TMM has an

opposite phenotype in stems; here, it results in a lack of stomata

(Geisler et al., 1998; Bhave et al., 2009). TMM is expressed in the

epidermis of both leaves and stems and in both organs is restricted

1 These authors contributed equally to this work.2 Address correspondence to [email protected] author responsible for distribution of materials integral to thefindings presented in this article in accordance with the policy describedin the Instructions for Authors (www.plantcell.org) is: Dominique C.Bergmann ([email protected]).CSome figures in this article are displayed in color online but in blackand white in the print edition.WOnline version contains Web-only data.OAOpen Access articles can be viewed online without a subscription.www.plantcell.org/cgi/doi/10.1105/tpc.111.086637

The Plant Cell, Vol. 23: 2864–2879, August 2011, www.plantcell.org ã 2011 American Society of Plant Biologists. All rights reserved.

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to stomatal precursor cells and their sisters (Nadeau and Sack,

2002).

Whereas TMM and the ERf are involved in overlapping devel-

opmental processes, the biochemical bases of their activities are

likely to be different. All four proteins possess extracellular LRR

repeats, but TMM, unlike the ERf receptors, lacks an intracellular

kinase domain. By analogy with signaling pathways in the shoot

meristem, it was hypothesized that TMM could participate in

active signal transduction only though physical interactions with

proteins that possess such domains (Nadeau and Sack, 2002;

Shpak et al., 2005). Alternatively, TMMmight modulate receptor

kinase signaling by forming inactive associations with RLKs or

altering availability of an RLK’s ligand (Shpak et al., 2005; Abrash

and Bergmann, 2010).

Strong candidates to be ligands of the ERf and TMM receptors

are members of the EPIDERMAL PATTERNING FACTOR LIKE

(EPFL) family. The EPFLs are small proteins predicted (and in one

case, demonstrated) to be processed into ;45– to 75–amino

acid long secreted peptides with compact Cys knot structures

(Kondo et al., 2009; Sugano et al., 2009). Three members of the

gene family, EPF1, EPF2, and EPFL9/STOMAGEN, have clear

roles in stomatal development. EPF1 and EPF2 are expressed

specifically within subsets of stomatal lineage cells, and STO-

MAGEN is expressed in leaf mesophyll; all three depend largely

upon TMM for their ability to affect stomatal development (Hara

et al., 2007, 2009; Hunt and Gray, 2009; Kondo et al., 2009;

Sugano et al., 2009). A fourth member of the family, EPFL6/

CHALLAH (CHAL), however, exhibits different behaviors. CHAL

was initially identified in a screen for suppressors of tmm-1

tissue-specific phenotypes, and consistent with this genetic

relationship (and in contrast with other EPFLs), phenotypic

effects ofCHAL loss or overexpression are revealed or enhanced

in a tmm mutant background. CHAL does not need TMM to

signal and, in fact, appears to be inhibited by it (Abrash and

Bergmann, 2010).

The diverse behaviors of EPFL family members, when con-

sidered in combination with TMM and the ER family and quan-

tified using the discrete and sensitive output of stomatal

development, provide a system in which to design and test

models of signaling specificity during plant development. Here,

we identify phenotypes for two additional EPFL family members

that align them with ER functions and that uncover redundancy

among related ligands. Our data suggest that different EPFL

family members mediate different ERf roles, with CHAL-related

family members dedicated for growth regulatory processes and

EPF1/2-related members specialized for stomatal regulation.

Together, our results point to a bufferingmechanism that permits

simultaneous, yet compartmentalized and distinct, function of a

common receptor set during tissue patterning and organ growth.

RESULTS

CHAL Family Genes Are Involved in Developmental

Processes outside of the Stomatal Lineage

To understand the mechanisms underlying CHAL’s unique and

tissue-specific relationship to the ERf and TMM, we asked

whether other EPFL family genes might be acting redundantly

withCHAL, thereby obscuringCHAL’s true functions. Among the

11 members of the EPFL family, EPFL4 and EPFL5 share with

CHAL a highly conserved sequence of amino acids (17/18

identical) located at the C terminus of the protein (Figure 1A).

Because of this sequence similarity and functional similarities

later revealed by our studies, it became useful to distinguish this

subfamily of EPFLs as CHAL family (CHALf) ligands; we will refer

to the CHAL paralogs as CHALLAH-LIKE1 (CLL1; EPFL5/

At3g22820) andCLL2 (EPFL4/At4g14723). We obtained and char-

acterized T-DNAmutant lines for each gene; cll1-1 (SALK_005080)

bears a T-DNA insertion in the 59 untranslated region and produces

reduced transcript, whereas cll2-1 (SALK_071065) bears a T-DNA

insertion in the second exon and appears to be a null allele

(Figures 1B to 1E). Neither single mutant displayed obvious

phenotypes related to stomatal development (data not shown;

Hara et al., 2009). The structural similarities among the three

CHALf members might lead to redundancy, such that a pheno-

type would be revealed only in multiple mutants; however, even

in chal cll1 cll2 triple homozygotes, stomatal pattern appeared

normal, and we observed no change in stomatal density (Fig-

ure 1F).

These results do not support a role for CHAL or its related

genes as stomatal development regulators in a wild-type back-

ground. While constructing multiple CHALf mutants, however,

we observed that a fraction of plants in segregating populations

displayed distinctive alterations in growth. Given that ER func-

tions in growth as well as stomatal patterning (Shpak et al., 2004,

2005) and that EPFLs were previously shown to act through the

ER family (Hara et al., 2007, 2009; Hunt and Gray, 2009), we

conducted a more comprehensive analysis of growth and de-

velopment in single, double, and triple mutant CHALf lines. We

found that plants bearingmutations in bothCHAL andCLL2 (chal

cll2 and chal cll1 cll2) are defective in growth, as evident from

their short stature, shortened pedicels, and compact inflores-

cences (Figures 1G to 1K). Several CHALf mutant lines also

display varying degrees of sterility, with chal cll1 cll2 producing

mostly sterile siliques (Figure 1G, inset). We quantified plant

height, pedicel length, and internode length in 7-week-old plants,

comparing CHALf mutants to both wild-type and er-105 (null)

mutants. In each of these characters, chal cll2 and chal cll1 cll2

mutants aligned with er, and both ligand and receptor mutants

were statistically different from the wild type (Figures 1L to 1N).

Having revealed new phenotypes for EPFL family members

outside of the stomatal lineage, wemeasured height and pedicel

and internode length in plants deficient for EPF1, EPF2, and

STOMAGEN. For none of these other EPFL genes, however, did

mutation or RNA interference (RNAi) knockdown produce con-

sistent growth differences from wild-type controls (see Supple-

mental Figure 1 online).

The shortened pedicels and stems of chal cll2 and chal cll1 cll2

could be due to a defect in cell expansion or, like er, in cell

proliferation (Shpak et al., 2004). To distinguish between these

possibilities, we examined the cortex cells of pedicels using

differential interference contrast (DIC) microscopy. We quanti-

fied the longitudinal length of adjacent cells in at least eight

pedicels per genotype. The average longitudinal length of 10

cells from the wild type was 226.63 6 33.5 mm; for er, 228.09 6

Specificity in Receptor–Ligand Interactions 2865

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Figure 1. Mutations in CHAL and Paralogs CLL1 and CLL2 Generate Nonstomatal Growth Phenotypes Resembling er Mutants.

(A) ClustalW2 (Chenna et al., 2003) alignment of C-terminal region of EPF-like proteins. The box indicates the region highly conserved between CHAL,

CLL1, and CLL2; asterisk, identical residue; colon, conserved substitution; dot, semiconserved substitution. Numbers immediately preceding and

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41.78 mm; and for chal cll1 cll2, 198.436 34.07 mm (mean6 SD).

Although chal cll1 cll2 cortex cells were slightly shorter in length

than the wild type (nonparametric multiple comparisons by si-

multaneous test procedure (STP), significant at a = 0.05 but not

at a = 0.025), this difference is not sufficient to account for a

reduction of pedicel length from 8.9 6 0.88 mm (wild type) to

3.13 6 0.39 mm (chal cll1 cll2, mean 6 SD). We therefore

conclude that, similar to er, the shortened pedicels of chal cll2

arise primarily through a defect in cell proliferation.

CHALf Genes Have Overlapping yet Distinct

Expression Patterns

Receptor–ligand interactions require common temporal expres-

sion and overlapping (or adjacent) spatial expression. We cre-

ated transcriptional reporters for CLL1 and CLL2 using 2.6 to 2.7

kb of 59 sequence driving b-glucuronidase (GUS) and assayed

GUS activity among progeny of at least four independent trans-

formed lines alongside previously published CHAL (Abrash and

Bergmann, 2010) and ERf transcriptional reporters (Shpak et al.,

2004) at various developmental stages to determine the extent of

their spatiotemporal overlap (Figure 2).

CHAL was initially characterized for its effects on hypocotyls,

and, consistent with previous reports (Abrash and Bergmann,

2010), we foundCHALwas expressed primarily in interior tissues

of this organ. In this study, we observed high expression at 36 h

after germination (Figure 2A) that ceased by 3 d after germination

(DAG) (Figure 2G). In three independent lines, CLL1 was also

observed in the hypocotyl, but its pattern was distinct from

CHAL; its expression was asymmetric and observed on the side

proximal to the folded cotyledons at germination (Figure 2B).

CLL2 expression was not observed in the hypocotyl (Figure 2C)

but was first seen at the base of the apical meristem at 3 DAG

(Figure 2I). When compared with ERf expression patterns, CLL2

is coincident with ERL2 expression (Figure 2L), and CHAL and

CLL1 expression in the young hypocotyl coincides with ER and

ERL1 expression (Figures 2D and 2E).

Given their mutant phenotypes in plant growth, we anticipated

that CHAL and CLL2 would be expressed in developing stems.

We observed strong expression of both CHAL and CLL2 in

developing stems soon after bolting (in plants <3 cm in height;

Figures 2M and 2O). In older plants, 10 to 20 cm in height, the

strongest expressionwas observed near the apex and in nascent

branches (Figures 2S and 2U), suggesting that expression within

an organ decreases as the organ ages and cells exit proliferative

stages. CLL1, by contrast, was generally not expressed at high

levels in stems (Figures 2N and 2T). These expression patterns

are consistent with publicly available tiling array data on the

CHALf genes (see Supplemental Figure 2J’ online). ERf genes

(Figures 2P to 2R and 2V to 2X) were also expressed in stems and

pedicels to varying degrees, with ERL2 showing the weakest

expression. ERf expression noticeably overlapped with CHALf

expression (Figures 2O, 2Q, 2S, and 2V).

CHALf genes are also expressed in the embryonic hypocotyl

(CHAL and CLL1; see Supplemental Figures 2D and 2E online),

the base of flowers (CLL1; see Supplemental Figure 2Q online),

young siliques (CLL2, two lines; see Supplemental Figure 2R

online), the chalazal region of ovules (CHAL and CLL1; see

Supplemental Figures 2J and 2K online), and near the root apex

(CHAL and CLL1; see Supplemental Figures 2V and 2W online).

We confirmed earlier reports of ERf expression in embryos and

ovule chalazal regions (Pillitteri et al., 2007) (see Supplemental

Figures 2G and 2H online) and also observed ERf expression in

the root (see Supplemental Figures 2S and 2U online). Previous

studies implicated ERf in integument proliferation, defects in

which lead to gametophyte abortion in erf mutants (Pillitteri et al.,

2007). The strong ovule expression of CLL1 (and to a lesser

extent, ofCHAL) may explain the sterility observed whenCLL1 is

removed in combination with other CHALf genes (Figure 1G,

inset). The GUS expression patterns are consistent with the

Figure 1. (continued).

following the sequence indicate the amino acid position within each respective protein.

(B) and (C) Schematics of CLL1 and CLL2 loci, respectively. Empty boxes, annotated untranslated regions; filled boxes, exons; lines, introns; triangles,

positions of T-DNA insertions.

(D) and (E) RT-PCR analysis of cll1-1 and cll2-1 alleles, respectively (ACT is ACTIN control; “Genomic” indicates genomic DNA template).

(F) Quantification of stomatal density in 8 DAG adaxial cotyledons. Data are displayed as boxplot graphs following standard conventions. Bold lines

indicate medians, boxes indicate quartiles above and below median, whiskers extend to most extreme value no more than 1.5 interquartile ranges from

box, and circles indicate values beyond whiskers. n$ 14 plants per genotype. ns, not significantly different from the wild type by Wilcoxon two-sample

test (P > 0.2).

(G) Aerial portions of 48 DAG Col, er-105, chal cll2, and chal cll1 cll2 plants. Inset of primary inflorescences shows sterility of chal cll1 cll2, and asterisks

mark sterile siliques. Bar = 5 cm.

(H) to (K) Inflorescence stem apices ;1 week after bolting illustrating reduction in pedicel elongation in er-105 (I), chal cll2 (J), and chal cll1 cll2 (K)

relative to Col (H).

(L) Quantification of plant height at 48 DAG; n = 10 plants per genotype.

(M) Quantification of pedicel lengths at 6.5 weeks of age; n = 50 pedicels per genotype, as five pedicels sampled from each of 10 individuals.

(N) Quantification of internode lengths at 6.5 weeks of age; n = 10 individuals per genotype. Values represent average internode length calculated over

four adjacent internodes.

In (L) to (N), letters a to d indicate maximal nonsignificant sets (MNS) to which the genotype belongs. Genotypes found in the same MNS are not

significantly different from one another, whereas genotypes not found in a common MNS are significantly different from one another, at the specified

significance level (a = 0.01 by nonparametric multiple comparisons by STP).

[See online article for color version of this figure.]


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Figure 2. Transcriptional Reporters for CHALf and ERf Are Coexpressed in Tissues Consistent with Their Loss-of-Function Phenotypes.

All images are of reporters based on promoters of indicated genes fused to GUS (Pro:GUS) and are incubated for times described in Methods.

(A) to (F) Pro:GUS expression at 36 h after germination; seedlings are oriented to provide view of hypocotyl, cotyledon bases, and tips of first leaves and

to capture the strongest expression levels. Specific reporters are indicated above each image, and each column represents the same reporter

construct.

(G) to (L) Pro:GUS expression at 3 DAG.

(M) to (R) Pro:GUS expression of primary inflorescence soon after bolting (plant height < 3 cm). Boxes mark nonstaining leaf in CLL2 (O) and staining

leaf in ERL2 (R).

(S) to (X) Pro:GUS expression in older primary inflorescence (plant height 10 to 20 cm).

Bars = 200 mm in (A) and (G) and 2 mm in (M) and (S). All images in a row are at the same magnification.

2868 The Plant Cell

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CHALf genes mediating a subset of the ERf’s diverse growth

functions, though notably, unlike the ERf (and other EPFLs),

CHALf genes do not appear to be expressed in cotyledons or

cauline leaves (Figures 2M to 2O and 2S to 2U; see Supplemental

Figures 2Y to 2B’ online). Examination of stomatal precursors

and guard cells indicates that, unlike ERfs, CHALf genes do not

exhibit elevated stomatal lineage expression (see Supplemental

Figures 2C’ to 2I’ online).

CLL1 Can Compensate for CLL2Growth Activity

CLL1 andCLL2 are very similar in sequence, but loss-of-function

and GUS expression studies suggest that these two proteins

have different endogenous roles. To determine whether expres-

sion pattern or biochemical activity is the primary difference

between CLL1 and CLL2, we tested whether CLL1 promotes

growth when expressed under control of the CLL2 promoter

(CLL2pro:CLL1). CLL2pro:CLL1 in chal cll2 qualitatively resem-

bled the wild type (19/21 T1s), whereas a negative control

(CLL2pro:GFP [for green fluorescent protein]) did not. We con-

firmed rescue through quantification of plant height, pedicel

length, and internode length in 12 CLL2pro:CLL1 T1s (see

Supplemental Figures 1A to 1C online) and three CLL2pro:

CLL2 T1s (positive control; see Supplemental Table 1 online).

These results indicate that both CLL1 and CLL2 have growth-

promoting activity and that expressional differences likely ac-

count for the behavior of the paralogs. They further demonstrate

that the CLL2 promoter fragment used in the GUS reporter

constructs is sufficient for rescue of the chal cll2 growth pheno-

type, confirming the validity of this reporter as a proxy for CLL2

expression.

CHAL Family Overexpression Phenotypes Are

ERf Dependent

The above analyses indicate that CHALf genes can function as

growth regulators, likely acting through the ERf. However, the

CHALf proteins are similar in sequence to stomatal regulators

EPF1, EPF2, and STOMAGEN (Hara et al., 2009; Kondo et al.,

2009), and CHAL itself was originally identified in a stomatal

screen (Abrash and Bergmann, 2010). In overexpression assays,

CHAL confers distinct, quantifiable stomatal phenotypes (Abrash

and Bergmann, 2010), and Hara et al. (2009) report in an EFPL

family survey that CLL1 and CLL2 can similarly repress stomatal

production. We therefore used the stomatal phenotypes of

overexpressors to measure CHALf activity and its mediation

by the ERf genes (and TMM).

First, we confirmed that CLL1 and CLL2 can inhibit stomatal

production inmultiple independent transformants.We found that

35Spro:CLL1 and 35Spro:CLL2 markedly inhibit stomatal pro-

duction when overexpressed in a wild-type background (Figure

3A), whereas 35Spro:CHAL T1 seedlings do not consistently

differ from the wild type in their stomatal production (Figure 3A).

In this regard, CLL1 and CLL2 bear some resemblance to the

stomatal EPFL family members EPF1 and EPF2, which are also

thought to act through the ERf.

When overexpressed in a tmm background, however, CLL1

and CLL2 behave similarly to CHAL (and unlike EPF1 and EPF2).

Rather than losing their activity in tmm, all three CHALf genes

confer a striking reduction in stomatal density, such that coty-

ledons of T1 plants are typically devoid of stomata and display

few or no asymmetric cell divisions (Figure 3B; see Supplemental

Figures 3B, 3F, and 3J online). Thus, rather than mediating

CHALf signaling, TMM actually functions to reduce or dampen

this signaling.

Using the sensitized tmm background, we then conducted a

series of experiments to assess whether one or more ERf

members mediate CHALf activity and whether receptor speci-

ficity varies among family members. To do so, we overexpressed

CLL1 and CLL2 in tmm backgrounds with reduced ERf activity

(specifically, backgrounds in which two of the three ERf genes

weremutated). Loss of any pair of ERf members was sufficient to

noticeably mitigate the overexpression phenotypes of CLL1 and

CLL2 in tmm, but the type and degree of phenotypic mitigation

was dependent upon the specific receptor remaining (Figure 3C;

see Supplemental Figure 3 online). In some backgrounds, only

asymmetric division was extensively rescued, whereas in other

backgrounds, differentiation was also considerably rescued. To

represent asymmetric division phenotypes, we show images

of typical cell division patterns for each genotype and present

a table of observed phenotypes (see Supplemental Figure 3

online). To capture the range of differentiation phenotypes ob-

served, we scored T1s by placing them into categories corre-

sponding to the cotyledon regions in which they produced

stomata (most to least rescued: central, peripheral, hydathode,

no stomata; Figure 3C). We observed the greatest rescue of

stomatal differentiation in tmm erl1 erl2mutants (active receptor:

ER), with a substantial fraction of CLL1 or CLL2 overexpressors

producing stomata in the periphery or center of the cotyledon

(Figure 3C; see Supplemental Figures 3E, 3I, and 3M online).

The phenotypes observed for CLL1 and CLL2 overexpression

in tmm backgrounds with ERf mutations are similar to those

observed for CHAL (Figure 3C; see Supplemental Figures 3B to

3E online; Abrash and Bergmann, 2010) and are consistent with

all three genes exerting their effects through the ERf. Although

CHAL, CLL1, and CLL2 overexpression phenotypes were not

identical, there were no clear-cut differences in receptor spec-

ificity among the three ligands, and, given the variability we have

observed in this assay, we do not think it can reveal subtle

differences in specificity. Similarly, though our findings show that

different ERf mutations modify CHALf overexpression pheno-

types in distinct ways, these differences may reflect the non-

identical endogenous activities of the receptor genes (e.g., ER

inhibits entry divisions, andERL1 andERL2 inhibit differentiation;

Shpak et al., 2005) rather than differences in their capacity to be

activated by the CHALf. Thus, our data suggest that CHALf

members can likely act through all three members of the ERf but

do not allow us to assign specific receptor-ligand pairings.

CHAL Family Mutations Affect Stomatal Production in

tmm Hypocotyls

Based on overexpression assays, CHALf members appear able

to activate signaling through the ERf receptors involved in

stomatal production. So why doesn’t endogenous CHALf ac-

tivity seem to influence stomatal development? Based on the


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expression patterns ofCHALf reporters, a simple lack of expres-

sion can account for the absence of phenotypes in many tissues,

including cotyledons. Then why aren’t clear stomatal pheno-

types observed in organs with CHALf expression, such as the

hypocotyl? Two basic possibilities come to mind: (1) CHALf

members never reach the stomatal lineage, due tomistimed and/

or subepidermal expression or (2) stomatal lineage-specific

factors actively block CHALf effects. The latter possibility was

of particular interest given that TMM, encoding a stomatal

lineage-specific coreceptor, seemed to strongly dampen CHALf

effects in overexpression assays (Figure 3).

If part of TMM’s role in the endogenous system is, in fact, to

exclude CHALf signaling from the stomatal lineage, then two

predictions should be fulfilled. First, reduction of CHALf activity

should have little impact on stomatal patterning in a wild-type

(TMM+) background because, if TMM prevents CHALf molecules

fromaffecting the epidermis, it would simply have nothing to block

when they were absent. Second, loss of TMM should enable the

CHALf genes to influence stomatal production, resulting in fewer

stomata where these genes are highly expressed.

To test these predictions, and to assess whether endogenous

CHALf members can reach and affect the stomatal lineage, we

examined CHALf mutant stomatal phenotypes in the presence

and absence of TMM. We scored phenotypes in hypocotyls

because the original chal phenotype was revealed in this organ

(Abrash and Bergmann, 2010), CHAL and CLL1 are expressed

here, and the number of stomata produced in the entire organ

can be evaluated. In a wild-type (TMM+) background, chal cll1

cll2 hypocotyls resembled the wild type in stomatal number and,

if anything, produced fewer stomata (perhaps due to growth

alterations; Figure 4A). In a tmm background, by contrast,

reduction of CHALf activity conferred a strong increase in hypo-

cotyl stomatal production. tmm single mutants produce virtually

no hypocotyl stomata, but tmm chal cll1 cll2 mutants produced

routinely over 100 per hypocotyl, often arranged in large clusters

(Figures 4B, 4C, and 4J). This result indicates that CHALf mem-

bers can indeed reach and affect the stomatal lineage when

present at their endogenous levels, in their endogenous spatio-

temporal patterns.

Based on analysis of single and multiple CHALf mutants in

tmm, CHAL and CLL1 appear primarily responsible for repres-

sion of stomatal production in tmm hypocotyls, consistent with

their expression in this organ (Figures 2A, 2B, and 4B to 4J).CLL2

may have some effect, but a consistent pattern was difficult to

discern due to high overall variability in phenotypic expressivity

and penetrance. To capture this variability as faithfully as pos-

sible, and because no single replicate was adequately repre-

sentative of our observations, we chose to show two replicates

for each genotype (Figure 4B).

CHAL Family Activity Explains a Subset of tmm Phenotypes

The suite of distinct and sometimes opposing tissue-specific

tmm phenotypes was recognized many years ago, but its un-

derlying cause has been elusive (Geisler et al., 1998). Our finding

that loss of TMM allows the CHALf genes to inappropriately

repress stomatal production in hypocotyls provides a plausible

explanation for phenotypes in this organ. Because CHALf genes

Figure 3. The Dependence of CHALf Overexpression Phenotypes on Receptor Genotypes Is Revealed through Stomatal Phenotype Assays.

(A) to (B) Quantification of effects of CHALf overexpression on stomatal density in wild-type and tmm adaxial cotyledons, respectively (10 DAG T1s).

Control = transformation with empty vector. Bold lines indicate medians, boxes indicate quartiles above and below median, and whiskers extend to

most extreme value no more than 1.5 interquartile ranges from box. n = 15 T1s per genotype in (A); n = 14 T1s per genotype in (B). *, significant at a =

0.05; **, significant at a = 0.01; ***, significant at a = 0.001 by nonparametric multiple comparisons by STP.

(C) Phenotypes of CHALf-overexpressing T1s in tmm and ERf mutant backgrounds (10 DAG). Each T1 was scored for rescue of stomatal differentiation

(for details, see Methods). From least to most rescued, categories were defined as the following: no stomata = no stomata in cotyledons; hydathode =

one or more stomata in hydathode region of cotyledon; peripheral = one or more stomata in cotyledon periphery; central = one or more stomata in

central cotyledon. The plants scored quantitatively in (B) are a subset of those scored qualitatively in (C).

2870 The Plant Cell

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are also expressed outside of hypocotyls, we asked whether

CHALf activity contributes to tmm stomatal underproduction phe-

notypes in otherorgans. In stems,muchas inhypocotyls, tmmchal

cll1 cll2mutants produced large numbers of stomata, dramatically

more than did tmm (Figure 4K). Thenumber of stomata in tmmchal

cll1 cll2 stems was considerably greater than in the wild type (see

Supplemental Figure 4 online), suggesting that loss of CHALf

function does not restore wild-type stem stomatal patterning but

rather causes stems to resemble other tmm organs that overpro-

duce stomata (e.g., rosette leaves). In adaxial sepals, however,

only a weak increase in stomatal density was observed upon loss

of CHALf activity, such that tmm chal cll1 cll2 had increased

stomatal number relative to tmm but still bore far fewer stomata

than the wild type (Figure 4N; data not shown). tmm chal cll1 cll2

sepals also appeared larger than tmm sepals, potentially contrib-

uting to the observed difference.

Figure 4. The Dependence of CHALf Loss-of-Function Phenotypes on TMM Is Revealed through Stomatal Phenotype Assays.

(A) Quantification of hypocotyl stomata in Col and chal cll1 cll2 (12 DAG, n = 20 plants per genotype). Bold lines indicate medians, boxes indicate

quartiles above and below median, and whiskers extend to most extreme value no more than 1.5 interquartile ranges from box. ns, not significantly

different from the wild type by Welch two-sample t test (P > 0.2).

(B) Quantification of hypocotyl stomata in CHALf mutants combined with tmm (12 DAG, n $ 10 plants per genotype). Each shaded or unshaded block

denotes a genotype, while letters denote replicates. Plants were grown in two replicate batches A and B, each of which included all eight genotypes.

(C) to (J) DIC images of 12 DAG hypocotyls with genotypes indicated in each panel.

(K) Quantification of inflorescence stem stomatal density (n $ 9 plants per genotype). ***, P < 0.001 by Welch two-sample t test.

(L) and (M) DIC images of inflorescence stems with genotypes indicated in each panel.

(N) Quantification of stomata in adaxial sepals (n $ 10 sepals per genotype). ***P < 0.001 by Wilcoxon two-sample test.

(O) Extent of stomatal clustering in adaxial cotyledons (8 DAG, n$ 20 plants per genotype). Clusters are categorized by the number of stomata (“mers”)

they contain; unclustered stomata are 1-mers. Shaded bars indicate mean; error bars are 6 SD.

Bars = 50 mm in (C) and (L); (C) to (J) are at the same scale, and (L) and (M) are at the same scale.


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We also examined the effects of reduced CHALf activity on an

organ in which tmm plants show increased stomatal density, the

adaxial cotyledon. We have not detected measureable CHALf

expression in this organ (Figures 2A to 2C; see Supplemental

Figures 2Z to 2B’ online), and in accord with this, did not detect a

consistent difference in density between tmm and tmm chal cll1

cll2 (see Supplemental Table 2 online). We did consistently

observe a shift in the size distribution of stomatal clusters in tmm

chal cll1 cll2, such that the fraction of stomata contacting other

stomata was higher than in tmm alone (0.377 6 0.106 in tmm

versus 0.5546 0.112 in tmm chal cll1 cll2, mean6 SD; P < 0.001

by Wilcoxon two-sample test; Figure 4O). As was noticed in the

sepals, in general, the tmm lines appeared less healthy than the

tmm chal cll1 cll2 lines, and this could indirectly affect some

measurements. Ultimately, it appears that CHALf activity can

largely explain loss of stomata from tmm stems and may con-

tribute to stomatal reduction phenotypes in sepals but, as

expected from expression patterns, does not contribute sub-

stantially to phenotypes in cotyledons.

Testing a Model for CHAL Family and EPF1/2 Interactions

with TMM

Our loss-of-function analyses suggest a model in which TMM

expression prevents stomatal lineage cells from perceiving

growth-regulatory CHALf signals. At the same time, TMM also

permits or enhances perception of EPF1 and EPF2 signals, which

are produced in stomatal precursors and enforce stomatal pattern

(Hara et al., 2007, 2009; Hunt and Gray, 2009). The relationship of

TMM to these two classes of genes, together with the expression

of TMM exclusively in the stomatal lineage, might permit efficient

segregation of growth- and stomatal-regulatory signaling despite

the spatial overlap of the respective ligands. In thismodel, two key

factors (interactions with TMM and expression patterns) distin-

guish growth- and stomatal-regulatory ligand classes. If these

factors were reversed, could these ligands substitute for one

another? We addressed this question by rewiring the stomatal

system to use growth-regulatory ligands. To do so, we removed

TMM from the epidermis, expressed CHAL under an EPF1 pro-

moter (EPF1pro:CHAL), and assayed CHAL’s ability to perform

EPF1’s endogenous role of enforcing stomatal spacing.

We transformedEPF1pro:CHAL intowild-type, epf1, tmm, and

tmm epf1 backgrounds. As previously reported, the EPF1 pro-

moter drives strong yellow fluorescent protein expression in

stomatal precursors and young stomata (Hara et al., 2007; see

Supplemental Figure 5 online) accompanied byweak, ubiquitous

fluorescence (see Supplemental Figure 5 online). As predicted by

our model, EPF1pro:CHAL did not decrease stomatal density or

reduce stomatal clustering in backgrounds with functional TMM

(wild type Columbia [Col] and epf1; Figures 5B and 5C, and 5H

and 5I versus 5D and 5E). In backgrounds lacking TMM activity

(tmm and tmmepf1), however, EPF1pro:CHAL conferred striking

alterations in stomatal density and spacing (Figures 5B and 5C,

and 5J and 5K versus 5F and 5G). Median stomatal density was

reduced to roughly wild-type levels, and the clustering profile

was strongly shifted toward single stomata and small clusters

(Figures 5B and 5C). Plants displaying a high degree of rescue

looked similar to the wild type in their overall pattern of differen-

tiated stomata, although small cells likely representing arrested,

transdifferentiated stomatal precursors were sometimes inter-

spersed with the stomata (Figures 5J and 5K, white arrows).

These results suggest that CHAL, when expressed in stomatal

precursors and in the absence of a TMM buffer, can indeed

function to enforce stomatal one-cell spacing.

DISCUSSION

Reuse of key signaling cascades in diverse processes and

contexts is a common theme in the development of multicellular

organisms. When multiple processes regulated by a single set of

receptors occur in close spatiotemporal proximity, signaling

specificity requires mechanisms that prevent spillover of activa-

tion from one process into another. The ERf receptors function in

a diverse range of developmental and physiological processes in

Arabidopsis, including growth regulation and patterning of sto-

mata. Here, we identified three EPFL family members, CHAL,

CLL1, andCLL2, as probable ERf agonists in regulation of growth.

Furthermore, we presented evidence that TMM, which encodes a

coreceptor expressed specifically in the stomatal lineage, acts to

block CHALf effects on stomatal development, enabling segre-

gation of the spatiotemporally overlapping, ERf-regulated pro-

cesses of growth and stomatal patterning. While we cannot

completely exclude the possibility that CHALf members act in

parallel to the ERf pathway, the hypothesis that CHALf members

act as growth-regulatory ligands for ERf receptors provides a

simple, unifying explanation for the mutant phenotypes and ge-

netic and physical interactions we observed.

Havingestablished thatCHALfmembers can functionasgrowth

regulators in pedicels and stems, we must reevaluate the rela-

tionship of these genes to stomatal development networks. Single

or higher order mutations in CHALf members can generate strong

stomatal phenotypes in tmm hypocotyls but fail to confer such

phenotypes in wild-type plants, indicating that TMM functions

endogenously to dampen or block CHALf signaling (in addition to

transducing appropriate stomatal-regulatory signals; see below).

As TMM expression is specific to, but widespread within, the

stomatal lineage, such a mechanism would effectively prevent

CHALf signals fromaffecting stomatal development. This in itself is

an elegant solution to the problem of specificity. By producing a

regulatory cofactor (TMM) in a spatially restricted domain (the

stomatal lineage), the plant prevents ligands appropriate to one

developmental process (growth) from interfering with a spa-

tiotemporally overlapping process (stomatal development) that

shares the same receptors.

When previous work on stomatal-regulatory EPFL family

members is considered, TMM emerges as an even more inge-

nious and economical innovation. Not only does this coreceptor

dampen growth-regulatoryCHALf signals, but it also potentiates

(and, indeed, is largely necessary for) transduction of stomatal-

regulatory EPF1, EPF2, and STOMAGEN signals (Hara et al.,

2007, 2009; Hunt and Gray, 2009; Kondo et al., 2009; Sugano

et al., 2009). Thus, just as growth-regulatory signals will not

aberrantly affect epidermal development because TMM is in the

stomatal lineage, so stomatal-regulatory signals will not aber-

rantly affect growth because TMM is absent from the internal

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tissues. The relationship of TMM to these two classes of ligands,

together with the expression of TMM exclusively in the stomatal

lineage, could thus permit the efficient segregation of growth-

and stomatal-regulatory signaling despite the spatial overlap of

the respective ligands (Figure 6).

Among animal signaling pathways, it is possible to find exam-

ples in which cofactors modulate ligand activity in a manner

similar to TMM. For instance, the glycosylphosphatidylinositol-

anchored protein Cripto plays a seemingly TMM-like role in the

interaction between the activin type II receptor and two classes

Figure 5. CHAL Can Enforce One-Cell Spacing When Expressed under the EPF1 Promoter in a tmm Background.

(A) Neighbor-joining tree depicting relationships between EPFL family proteins (modified from Rowe and Bergmann, 2010). Shading indicates proteins

of known function.

(B)Quantification of the effects of EPF1pro:CHAL on stomatal density (versus empty vector) in 10 DAG abaxial cotyledons. Bold lines indicate medians,

boxes indicate quartiles above and below median, and whiskers extend to most extreme value no more than 1.5 interquartile ranges from box. n = 15

T1s for each line. ns, not significant (P > 0.30 by Wilcoxon two-sample test); ***, P < 0.001 by Wilcoxon two-sample test.

(C) Quantification of the effects of EPF1pro:CHAL on stomatal clustering (versus empty vector) in 10 DAG abaxial cotyledons. Shaded bars indicate

mean; error bars are 6 SD.

(D) to (K) Representative DIC images of EPF1pro:CHAL and empty vector T1s. Genotypes and transgenes are indicated on each panel. White brackets

indicate clustered stomata; white arrows indicate apparent transdifferentiated cells observed in strongly rescued lines.

Bar = 50 mm in (D); (D) to (K) are at the same scale.

[See online article for color version of this figure.]


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of ligands in the TGF-b superfamily: it is required for Nodal-class

ligands to signal through this receptor but impedes the ability of

activin-class ligands to do the same (Kirkbride et al., 2005; Kelber

et al., 2008). As these relationships have been worked out largely

in tissue culture, it would be interesting to examine whether

Cripto has an in vivo role similar to that of TMM (i.e., whether it

functions to segregate activin II–dependent signaling pathways

that operate in close proximity).

Our findings that characterized EPFL family members cluster

into process-related subgroups raises the question of what roles

the remaining EPFL genes (Figure 5A) might play. The five

uncharacterized EPFL family members could encode stomatal

regulators, growth regulators, or ligands active in some other

ERf-dependent process, such as bacterial or fungal pathogen

resistance (Godiard et al., 2003; Llorente et al., 2005). In neither

growth nor stomatal development does loss of the currently

characterized EPFLs fully recapitulate the suite of defects gen-

erated by loss of the ERf. chal cll1 cll2 triple mutants do not have

the blunted siliques and altered leaf morphology characteristic of

er mutants and are clearly not as dwarfed as er erl1 erl2. This

suggests that additional EPFL genes may regulate growth and

may do so redundantly with the CHALf. In support of this

possibility, we found that two more EPFL family members,

EPFL1 (At5g10310) and EPFL2 (At4g37810), display expression

patterns roughly similar to the CHALf in publicly available tiling

array data (see Supplemental Figure 2K’ online; Laubinger et al.,

2008) and display CHAL-like behaviors in overexpression as-

says: No obvious stomatal phenotypes were reported for these

genes in a wild-type background (Hara et al., 2009), but we find

that they noticeably reduce stomatal density in tmm (see Sup-

plemental Figure 6 online). Activity of these genes might also

account for tmm phenotypes not explained by CHALf activity,

such as loss of stomata from the adaxial sepal.

Activity of additional EFPLs could also contribute to the

substantial phenotypic variability seen for CHALf mutants in a

tmm background. Despite care in standardizing growth condi-

tions and seedling selection, we observed considerable vari-

ability in stomatal phenotypes among seedlings of the same

genotype (Figure 4B). Such variability could be explained if

expression of CHALf members (and of spatially overlapping,

TMM-inhibited EPFLs) were constrained within a range appro-

priate to growth regulation but not conducive to consistent levels

of stomatal production in the absence of TMM. This hypothesis

might also account for the basal phenotypic variability observed

in tmm single mutants. Alternatively, this variability might reflect

the capacity of ERf receptors and/or their downstream mitogen-

activated protein kinase cascade to transduce environmental

cues and stresses (Godiard et al., 2003; Llorente et al., 2005;

Nakagami et al., 2005), perhaps coupled with enhanced sensi-

tivity to such cues and stresses in the absence of TMM. Inde-

pendent of its cause, this variability must be considered when

Figure 6. Models in Which CHALf Acting through ERf Receptors in a

Pathway Inhibited by TMM Can Explain Observed Phenotypes.

(A) Model for CHALf and EPF1/2 interactions with TMM and ERf

receptors in different regions of wild-type and tmm seedlings. Arrows

represent positive influences on signaling or developmental processes;

T-bars represent negative influences. Grayed-out elements are inactive

or not present in a particular organ or genotype. The positive regulator of

stomatal development EPFL9/STOMAGEN also requires TMM for activ-

ity, but for simplicity, is not shown here. The left half of the seedling

depicts the wild type (TMM+). In both the cotyledon and hypocotyl,

stomatal lineage ligands EPF1/2 signal through ERf and TMM receptors

to inhibit stomatal development for proper patterning. CHALf ligands in

the hypocotyl do not influence stomatal patterning due to the presence

of TMM, which dampens CHALf signaling. The right half of the seedling

depicts the tmm mutant: Without TMM, EPF1/2 do not inhibit stomatal

development, leading to the formation of stomatal clusters in cotyledons.

The opposite effect is observed in the tmm hypocotyl, however, due to

the presence of CHALf ligands in this organ. Without TMM’s dampening

effect, CHALf ligands overactivate ERf receptors, resulting in a strong

inhibition of stomatal development.

(B) Model for TMM compartmentalization of ERf receptor functions in

growth and stomatal development in stems. Apoplast-mobile CHALf and

EPF1/2 ligands can encounter ERf receptors both in the stomatal lineage

and in nonstomatal cells, such as the inner tissues. TMM, which is

present only in the stomatal lineage, differentially regulates these two

ligand classes. Thus, CHALf signaling does not affect stomatal devel-

opment because it is dampened by TMM, while EPF1/2 signaling does

not affect growth because it is potentiated by TMM. Arrows represent

positive influences on signaling pathways or developmental processes;

T-bars represent negative influences. Grayed-out elements represent

inactive pathways.

2874 The Plant Cell

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evaluating published experiments and designing new ones with

EPFL genes in a tmm background.

Characterization of the diverse functions of EPFL family mem-

bers represents one challenge for the future. Another such

challenge will be to clarify the biochemical interactions among

EPFL family ligands, ERf receptors, and TMM and to identify

downstream outputs to measure the signaling capacities of

specific ligand and receptor complexes. Genetic and phenotypic

evidence suggests that EPFLs act through the ERf; we would

therefore expect the ligands to physically interact with the

extracellular domains of ERf receptors. TMM’s dampening effect

on CHALf signaling, however, could arise via several different

mechanisms. In one scenario, TMM might bind CHALf mem-

bers directly, either titrating them away from the ERf or en-

gaging them in unproductive ERf/TMM/CHALf complexes.

Alternatively, TMM might not bind CHALf ligands directly, but

instead tie up ERf receptors in TMM/ERf complexes incapable

of CHALf binding. Rigorous demonstration of the mode of TMM

activity will require purification of endogenous complexes from

relevant tissues and awaits future studies. Binding of CHALf

members to specific ER and TMM extracellular fragments can

be observed in yeast two-hybrid assays (see Supplemental

Figure 7 online), consistent with models based on ligand

titration or formation of unproductive ERf/TMM/CHALf com-

plexes. In stomatal lineage cells, the abundance of receptors

and their differential affinities for ligands may further constrain

system behavior. Modeling approaches such as those recently

used to evaluate alternative models for shoot meristem LRR-

RLK–mediated signaling (Sahlin et al., 2011) will also be im-

portant hypothesis testing tools, especially as the number of

potential ligand and receptor players increases beyond what

can be easily tested empirically.

The function of TMM as a pathway buffering/signaling

specificity element represents, thus far, a unique role for an

RLP in plant LRR-RLK signaling. What is the potential for this to

be a widespread phenomenon? Among the;220 Arabidopsis

LRR-RLKs (Shiu and Bleecker, 2001) and 57 LRR-RLPs (Wang

et al., 2008), only two developmental partnerships have been

described: that of the ERf and TMM in growth and stomatal

development and that of RLK CLAVATA1 (CLV1) and RLP

CLV2 in shoot meristem maintenance. The latter appears to

follow fundamentally different rules than the former: CLV2

likely forms an active signaling complex with a unique in-

tracellular kinase (CORYNE; Muller et al., 2008), may signal

largely in parallel to CLV1 (Nimchuk et al., 2011), and is

expressed in a broader domain than is CLV1. Thus, it is unlikely

that RLPs as a category are dominated by TMM-like behaviors.

Rather, such coreceptor behaviors might be expected in cases

where: (1) a core set of receptors is used in multiple contexts

during the life of the organism, (2) the processes regulated

by the receptors occur in close spatiotemporal proximity, and

(3) ligands are mobile, such that those intended to act in one

process could bleed over into another. It will be intriguing

to see if other systems with these characteristics indeed

have TMM-like mechanisms for ligand discrimination and, if

not, even more intriguing to see what other paths evolution

has found to achieve compartmentalized signaling in close

quarters.

METHODS

Plant Materials and Growth Conditions

Col-0 or Col-0 containing the E1728 guard cell GFPmarker (Gardner et al.,

2009) was used as the wild type. Alleles used were cll1-1 (SALK_005080),

cll2-1 (SALK_071065), chal-2 (Abrash andBergmann, 2010),er-105, erl1-2,

and erl2-1 (Shpak et al., 2004), tmm-1 (Nadeau and Sack, 2002), epf1-1

(Hara et al., 2007), and epf2-1 (Hunt and Gray, 2009). The left border

junction of the cll1 T-DNA has sequence 59-AATTTGATTACTCTTT-

GACTTTGGACCACT-39, while the left border junction of the cll2 T-DNA

has sequence 59-TCAATTTGTTAGTAATAAGCGGTTTGGTGG-39 (under-

line indicates T-DNA; plain text indicates genomic DNA). With the excep-

tion of er-105, the allele number is dropped when these mutations are

referred to in the text. tmm er erl1, tmm er erl2, and tmm erl1 erl2mutants

(Shpak et al., 2005) were gifts of K.U. Torii. tmm er erl2 is also glabrous.

epf1 epf2 was a gift of J.E. Gray. STOMAGEN RNAi lines (Sugano et al.,

2009) were a gift of I. Hara-Nishimura. tmm epf1, multiple CHALf mutants,

and CHALf mutants in tmm backgrounds were generated by crossing and

isolated based on phenotypes and/or PCR genotyping (see below). tmm,

tmm epf1, and some CHALf mutant lines in tmm backgrounds contain the

E1728 guard cell GFP marker. ERpro:GUS, ERL1pro:GUS, and ERL2pro:

GUS lines (Shpak et al., 2004) were gifts of K.U. Torii. CHALpro:GUS lines

have been previously described (Abrash and Bergmann, 2010) and also

contain the E1728 guard cell GFP marker. Seeds were stratified at 48C

for$2 d. Seedlingswere grown on 0.5Murashige andSkoog (MS)medium

(Murashige and Skoog, 1962), 1% agar plates at 228C under constant light

in an incubator (I36LL; Percival). For later analyses, seedlings were trans-

ferred to soil and maintained in a growth chamber (AR66; Percival) at 226

18C with a 16-h-light/8-h-dark cycle.

Genotyping

Representative, functional primer pairs for genotyping (a subset of those

used over the course of the experiment) and sequences of gene-specific

primers are provided in Supplemental Table 3 online. T-DNA internal

primers LBa1 and LBb1.3 (Alonso et al., 2003; http://signal.salk.edu) and

JL-202 (Young et al., 2001) have been previously described.

RT-PCR

For analysis of cll1 and cll2, RNA was extracted from ;13 DAG whole

seedlings using the RNeasy plant mini kit (Qiagen) with on-column DNase

digestion. cDNA was synthesized with oligo(dT) primers using the Su-

perscript III first-strand cDNA synthesis kit (Invitrogen), followed by

amplification using Accuprime Pfx DNA polymerase (Invitrogen) and

gene-specific primers (see Supplemental Table 3 online) with program

958C for 2:00, 35 cycles of 958C for 0:15, 548C (forCLL2 andACT; 588C for

CLL1) for 0:30, 688C for 1:00, and final extension of 688C for 5:00. Bands

were visualized via ethidium bromide staining.

Plant Vector Construction and Transformation

Most manipulations involved Gateway-mediated cloning following the

manufacturer’s instructions. Primers used for each step are given in

Supplemental Table 3 online. For 35Spro:CLL1, the CLL1 coding se-

quence was amplified from clone U63004 (ABRC), cloned into pENTR/

dTOPO (Invitrogen), and recombined into pH35GS (Kubo et al., 2005)

using LR Clonase II (Invitrogen). For 35Spro:CLL2, cDNA clone

DQ487600 (ABRC) was recombined into pH35GS. For 35Spro:EPFL1,

clone GC63002 (ABRC) was recombined into pH35GS. For 35Spro:

EPFL2, the EPFL2 coding sequence was cloned into pENTR and

recombined into pH35GS. 35Spro:CHAL was described (Abrash and

Bergmann, 2010). The empty vector control for overexpression assays,


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pH35S, was generated from pH35GS by XbaI/XhoI excision of the

Gateway followed by blunting and self-ligation. For CLL1pro:GUS and

CLL2pro:GUS, 2.6- to 2.7-kb 59 regions were amplified from BAC

MIWI23 (ABRC) for CLL1 and BAC T2D4 (ABRC) for CLL2. Fragments

were cloned into pENTR/dTOPO and recombined into pBGGUS (Kubo

et al., 2005). For EPF1pro:YFP, the promoter region defined by Hara

et al. (2007) was cloned into pENTR/dTOPO and recombined into pHGY

(Kubo et al., 2005). For EPF1pro:CHAL, the same EPF1 promoter was

inserted in the NotI site of pENTR-CHAL (generated by amplifying the

CHAL cDNA out of clone DQ446581 [ABRC] and cloning into pENTR/

dTOPO) and the EPF1pro:CHAL fusion recombined into pHGS (a

pH35GS derivative generated by HindIII/XbaI excision of 35Spro fol-

lowed by blunting and self-ligation). The empty vector control for the

EPF1pro assay, pHS, was made from pH35GS by HindIII/XhoI excision

of 35Spro and the Gateway followed by blunting and self-ligation. For

CLL2pro:CLL2 and CLL2pro:CLL1, CLL2pro was excised from pENTR/

dTOPO using flanking NotI sites. CLL2 was amplified and cloned into

pENTR/dTOPO. CLL1 and CLL2 in pENTR/dTOPO were NotI digested,

and CLL2pro was inserted into the 59 NotI site. Constructs were then

recombined into pGWB1 (Nakagawa et al., 2007). CLL2pro:GFP was

generated by recombining CLL2pro pENTR/dTOPO into pGWB4

(Nakagawa et al., 2007).

Stably transgenic plants were generated byAgrobacterium tumefaciens–

mediated transformation using the floral dip protocol (Clough and Bent,

1998). Transgenic seedlings were selected either by growth on 1% agar +

0.5 MS plates supplemented with 50 mg/mL hygromycin (pH35GS, pHGY,

pGWB1, and pGWB4 based constructs) or in soil treated with Basta (Finale

1:2000 dilution; pBGGUS-based constructs).

Analysis of Stomatal Phenotypes

Preparation of Tissue

Multiple plant tissues were assayed for stomatal production. For hypo-

cotyls and cotyledons, plants were grown on 0.53 MS, 1% agar plates

until time of collection (noted for each experiment in the main text). For

analysis of inflorescence stem stomata, seedlings were moved from

plates to soil at 10 DAG (Figure 4K) or 11 DAG (see Supplemental Figure

4 online) and the third most basal internode of themain stemwas scored

at 37 DAG (Figure 4K) or 46 DAG (see Supplemental Figure 4 online). For

sepals, seedlings were moved from plates to soil at 11 DAG and scored

at 46 DAG. Sepals were collected from the oldest pedicel on the main

stem that still bore nonsenescent sepals.

Fixation and Mounting

Tissue samples were processed by one of three methods, depending on

tissue type and age: (1) Samples were collected into 70% ethanol,

cleared overnight at room temperature, and stored at 48Cas needed and

then rinsed in water and mounted in Hoyer’s medium (Liu and Meinke,

1998). For stem samples, an epidermal peel was taken from the cleared

sample following the water rinse. (2) Samples were cleared and stored in

7:1 ethanol:acetic acid at room temperature, treated with 1 N potassium

hydroxide (30 min to overnight), rinsed in water, and mounted in Hoyer’s

medium. (3) Fresh tissue was mounted directly in Hoyer’s medium and

placed in a vacuum chamber for ;1 week or imaged directly if epider-

mal peel. The second method was typically most effective in preserving

cell morphology.

Analysis and Scoring

Most images of stomatal phenotypes were obtained from samples

mounted in Hoyer’s medium and visualized using DIC microscopy on a

Leica DM2500 microscope. For quantitative analysis of hypocotyl and

adaxial sepal stomatal number, scoring was performed on the micro-

scope, and results reflect analysis of the entire organ or tissue. For

analysis of cotyledon stomatal density and clustering, and of inflores-

cence stem stomatal density, images were taken at 320 magnification

(0.320-mm2 field), and stomata and/or clusters were counted manually

from images using the Cell Counter in ImageJ (NIH). Only those

stomata/clusters contained fully within the frame of the image were

counted. For cotyledons, images were taken in the central region

overlying the distal vascular loop. If a cotyledon was too small to fill

the320 field, it was positioned so as to best fill the field and the stomata

within the frame were counted. For inflorescence stems, images were

randomly positioned within the regions of sample that were wide

enough to fill the camera field. Except in cases where differences

between genotypes were very obvious, images were randomized

before scoring and scored blind.

For qualitative (category) scoring of CHALf overexpressing T1s in tmm

and ERf mutant backgrounds, the adaxial and abaxial sides of all

cotyledons present on the plant were examined for stomata, and each

plant was placed into the “most rescued” category to which it belonged

(most to least rescued: central stomata, peripheral stomata, hydathode

stomata, no stomata). A single stoma in a given category was sufficient to

place a plant into that category. Stomata were scored as “central” if

contained within the vascular loops, “peripheral” if outside the vascular

loops but not in the hydathode, and “hydathode” if present at the distal tip

of the leaf, in or associated with the hydathode. Plants with stomata

placing them in a higher category (e.g., central) often, but not always, also

possessed stomata corresponding to lower categories (e.g., hydathode).

One to three batches of a given genotype + construct combination were

scored and pooled to give the final data presented.

Except in analyses of CHALf overexpressing T1s, which were often

unhealthy due to lack of stomata, only seedlings that met a minimum

health cutoff (based primarily on the presence and size of true leaves or on

the size of cotyledons) were included in assays.

Analysis of Growth Phenotypes

Plant growth was quantified at;7 weeks after germination. Plants were

grown on 0.53MS, 1% agar plates for 2 weeks and then transplanted to

soil. Height measurement was recorded as the distance from rosette to

the flower tips of the longest inflorescence. For chalf lines, the length of

five pedicels per plant (pedicels 3 to 7 from the base of the primary

inflorescence) wasmeasured. ForSTOMAGENRNAi lines, epf1 epf2, and

rescue/complementation experiments (CLL2pro:CLL1, CLL2pro:GFP,

and CLL2pro:CLL2 in chal cll2), the length of two pedicels per plant

(pedicels 6 and 7 from the base of the primary inflorescence) was

measured. For internodes, the distance between pedicels 3 and 7 for

each plant was recorded and then divided by 4 to determine a single

average internode length per plant. For chalf analysis, 10 plants were

analyzed per genotype for all measurements. For epf1 epf2 and STO-

MAGEN RNAi analysis, nine plants were analyzed per genotype/inde-

pendent line for all measurements (except STOMAGEN RNAi pedicels;

eight plants per linewere examined).MUTEpro:nucGFP (MacAlister et al.,

2007) in Col was used as a BASTA-resistant control to compare with

STOMAGENRNAi lines. For rescue of chal cll2 byCLL2pro:CLL1 (and the

CLL2pro:GFP control), 12 independent T1s were analyzed for height, and

11 independent T1s were analyzed for pedicel and internode length. For

complementation of chal cll2 by CLL2pro:CLL2, three independent T1s

were analyzed for each measurement.

To analyze cortex cell length, pedicels were analyzed at 6 to 7 weeks

using DIC at 340 on a Leica DM2500 microscope by focusing on the

subepidermal cortex plane. The length of a longitudinal cell file (10 cells

per cell file for data shown) was measured using ImageJ software (NIH).

Two measurements were taken per pedicel, and two pedicels of four

2876 The Plant Cell

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plants per genotype were analyzed, for a total of 16 measurements per

genotype. Two replicates were examined.

Statistical Analysis

Most statistical analyses were performed in R (http://www.r-project.

org/). For statistical analyses, each data point was treated as a

separate observation, even when multiple data points were obtained

from a single plant (e.g., in measurement of pedicel lengths). Data were

checked for normality using the Shapiro-Wilk test [shapiro.test() func-

tion]. If all samples in a given data set were normally distributed,

parametric analyses based on the normal distribution were used. For

comparison of two samples, the Welch two-sample t test [t.test()

function, unpaired, two-tailed] was used. If not all samples in a given set

were normally distributed [or, in the case of multiple comparisons, if

samples were normally distributed but differed significantly in variance by

Bartlett’s test using the bartlett.test() function], nonparametric analyses

were used instead. For comparison of two samples, the Wilcoxon two-

sample test [wilcox.test() function, unpaired, two-tailed] was used. The

reported P value is that returned by the function, but it should be noted

that this is not an exact P value when ties are present (as they typically

were). For multiple comparisons, nonparametric multiple comparisons

by STP were performed, as described by Sokal and Rohlf (1995).

Additional tables of critical values of the studentized range were

obtained from Harter (1960). Because nonparametric multiple compar-

isons by STP require equal sample sizes, samples were randomized

and then truncated to the size of the smallest sample included in the

analysis. As no R function is available for nonparametric multiple

comparisons by STP, a simple Java program was written to calculate

the test statistics, while cutoff values were calculated by hand. Test

statistics were not corrected for between-sample ties as the correction

is typically small and uncorrected values are conservative. a Values

indicate the experiment-wise error rate at which the comparison is

significant. When maximal nonsignificant sets were reported, they were

determined according to the definition of Rafter et al. (2003). Through-

out, the statistical method used for each comparison is noted in the

caption or text accompanying the data.

Boxplots were generated in R using the boxplot() function. The

boxplot is a standard graphical display format (Sokal and Rohlf, 1995)

in which a central horizontal line indicates the median, boxes above

and below the line indicate quartiles above and below the median, and

whiskers extend to the most extreme value no more than 1.5 interquartile

ranges from the box. Observations outside the whiskers (sometimes

called outliers) are depicted individually as circles. We treated these

points as outliers for purposes of graphical display only and did not

exclude them in statistical tests of significance.

Statistical comparisons between different CHALf mutant genotypes in

a tmm background were not performed for hypocotyl data due to the high

degree of within-genotype variation (as illustrated by the presentation of

data in Figure 4B).

Yeast Two-Hybrid Assays

Yeast two-hybrid analysis was performed with the matchmaker Two-

Hybrid System 3 (Clontech) using plasmids and conditions specified

by the manufacturer. cDNAs corresponding to ligands were PCR

amplified and cloned into pGBKT7 and receptor fragments into

pGADT7 (primers in Supplemental Table 3 online). TPD1 cDNA was

a gift of Dazhong Zhao (Jia et al., 2008). Pairwise interactions were

conducted in triplicate at 1:100 and 1:1000 dilutions after incubation

for 2, 3, and 4 d. Interactions were evaluated as positive if significant

growth (;50% saturation) was observed by day 3.

Accession Numbers

The Arabidopsis Genome Initiative identifiers for the genes mentioned in this

article are as follows: CHAL/EPFL6 (At2g30370), CLL1/EPFL5 (At3g22820),

CLL2/EPFL4 (At4g14723), EPF1 (At2g20875), EPF2 (At1g34245), EPFL1

(At5g10310),EPFL2 (At4g37810),STOMAGEN (At4g12970),ER (At2g26330),

ERL1 (At5g62230),ERL2 (At5g07180), TMM (At1g80080), TPD1 (At4g24972),

andMUTE (At3g06120).

Supplemental Data

The following materials are available in the online version of this article.

Supplemental Figure 1. Redundancy and Range of Nonstomatal

Phenotypes within the EPFL Family.

Supplemental Figure 2. Pro:GUS and Tiling Array Expression of the

CHALf and ERf in Additional Tissues.

Supplemental Figure 3. Qualitative Phenotypes of CHALf Over-

expression in Receptor Mutant Backgrounds.

Supplemental Figure 4. Stem Stomatal Density in tmm, tmm chal cll1

cll2, and Wild-Type Plants.

Supplemental Figure 5. Expression Pattern of EPF1pro:YFP.

Supplemental Figure 6. Overexpression Phenotypes of Additional

EPFL Family Members in tmm.

Supplemental Figure 7. Yeast Two-Hybrid Analysis of Interactions

between EPFL Ligands and Receptor Fragments.

Supplemental Table 1. CLL2pro:CLL2 Complementation of chal cll2.

Supplemental Table 2. Stomatal Density in tmm and tmm chal cll1

cll2 Adaxial Cotyledons.

Supplemental Table 3. Primers Used in This Work.

ACKNOWLEDGMENTS

We thank D. Zhao, I. Hara-Nishimura, K. Torii, and J. Gray for reagents,

Akhila Bettadapur for lab assistance, E. Shpak for insights, and mem-

bers of our lab for discussions and critical reading of the manuscript.

This work was supported by a Stanford Terman fellowship, by National

Science Foundation Grant IOS-0844521, and by a Stanford Vice Provost

for Undergraduate Education Major Grant to E.B.A. K.A.D. and E.B.A.

are National Science Foundation Graduate Research Fellows.

AUTHOR CONTRIBUTIONS

E.B.A., K.A.D., andD.C.B. designed research andwrote the article. E.B.A.

and K.A.D. performed research and analyzed data.

Received April 19, 2011; revised August 4, 2011; accepted August 15,

2011; published August 23, 2011.

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