{"paper_id":"0c2b52ea-e0f1-4fd6-bbb4-bc4f75f475db","body_text":"SPEECHLESS duplication in grasses expands potential for environmental regulation of stomatal development | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return\"[object Function]\"==o.call(a)}function e(a){return\"string\"==typeof a}function f(){}function g(a){return!a||\"loaded\"==a||\"complete\"==a||\"uninitialized\"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){(\"c\"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){\"img\"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),\"object\"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height=\"0\",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),\"img\"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||\"j\",e(a)?i(\"c\"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName(\"script\")[0],o={}.toString,p=[],q=0,r=\"MozAppearance\"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&\"[object Opera]\"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?\"object\":l?\"script\":\"img\",v=l?\"script\":u,w=Array.isArray||function(a){return\"[object Array]\"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split(\"!\"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split(\"=\"),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(\".\").pop().split(\"?\").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split(\"/\").pop().split(\"?\")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&\"css\"==i.url.split(\".\").pop().split(\"?\").shift()?\"c\":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results SPEECHLESS duplication in grasses expands potential for environmental regulation of stomatal development View ORCID Profile Joel M. Erberich , Britney Bennett , View ORCID Profile Dominique C. Bergmann doi: https://doi.org/10.1101/2025.07.29.667563 Joel M. Erberich 1 Department of Biology, Stanford University , Stanford, CA 94305, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Joel M. Erberich Britney Bennett 1 Department of Biology, Stanford University , Stanford, CA 94305, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Dominique C. Bergmann 1 Department of Biology, Stanford University , Stanford, CA 94305, USA 2 Howard Hughes Medical Institute , Stanford, CA 94305, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Dominique C. Bergmann For correspondence: dbergmann{at}stanford.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF Summary Plants regulate stomatal development and function to acquire atmospheric carbon dioxide for photosynthesis while minimizing water loss. The ancestral basic helix-loop-helix transcription factor (TF) gene that drove stomata production in early land plants diversified to become paralogs SPEECHLESS ( SPCH ), MUTE , and FAMA . Extant grasses exhibit a particularly interesting set of duplications and losses of SPCH . Using phylogenetic methods, we tracked the history of SPCH duplications. Brachypodium distachyon and Oryza sativa plants bearing mutations in either SPCH1 or SPCH2, and B. distachyon plants with SPCH1 or SPCH2 translational reporters were assayed under different environmental conditions for their effects on stomatal development. We identified the Poaceae-specific rho whole genome duplication as the origin of SPCH1 and SPCH2 and demonstrated that both paralogs remain under selection. We found paralog-specific divergence in response to two environmental perturbations in both B. distachyon and O. sativa . Plausible molecular mechanisms underpinning paralog divergence, and cellular mechanisms driving the stomatal phenotypes are supported by analyses of BdSPCH1 and BdSPCH2 RNA and protein expression and by sequence variation among grasses. These studies suggest ways in which a duplication of a key stomatal regulator enables adaptation and could inform genetic strategies to mitigate anticipated stressors in agronomically important plants. Introduction Stomata are cellular complexes that enable efficient gas exchange between the aerial portions of plants and the environment. Stomata appeared in the fossil record coincident with the appearance of land plants and, along with the waxy cuticle, are thought to enable plants’ success on land ( Payne, 1979 ). Among most clades of living and fossil plants, stomatal morphology is simple–two epidermal guard cells flanking a central pore. The morphology of neighboring cells and patterns of stomatal distribution on leaves vary considerably, however, and are useful taxonomic characters ( Bergmann and Sack, 2007 ). Based on anatomical observations of preserved samples, it has been hypothesized that the developmental pathways to create stomata have increased in complexity over evolutionary time ( Rudall et al ., 2013 ). In parallel, genes encoding the core fate-regulating transcription factors have also expanded. Phylogenetic studies link the origin, duplication and diversification of subfamily Ia and subfamily IIIb basic helix-loop-helix (bHLH) transcription factors (TFs) to stomatal lineage diversity ( Chen et al ., 2017 ; Clark et al ., 2022 ; Harris et al ., 2020 ; Pires and Dolan, 2010 ). Genetic studies in the bryophyte moss Physcomitrium patens found a single subfamily Ia and subfamily IIIb pair are required to produce stomata at the base of the sporophyte spore capsule where they serve to promote dehiscence ( Chater et al ., 2016 ). In the dicot angiosperm Arabidopsis thaliana , stomata are the end product of a multi-step developmental pathway. Initiation, commitment, and differentiation steps in this pathway are each uniquely regulated by a different stage-specific subfamily Ia bHLH, SPEECHLESS (SPCH), MUTE, or FAMA, respectively ( Bergmann et al ., 2004 ; Ohashi-Ito and Bergmann, 2006 ; Pillitteri et al ., 2008 ). Two Class IIIb factors, SCREAM (SCRM1) and SCRM2 are redundant heterodimer partners for all three subfamily Ia factors ( Kanaoka et al ., 2008 ). Orthologues of SPCH, MUTE, FAMA, SCRM1 and SCRM2 can be identified in many angiosperm genomes, and functional studies in Solanum lycopersicum (tomato), Oryza sativa (rice), Brachypodium distachyon (purple false brome) and Zea mays (maize) found these TFs to be generally conserved in their expression patterns and function in stomatal development ( Liu et al ., 2009 ; Nir et al ., 2023 ; Ortega et al ., 2019 ; Raissig et al ., 2016 ; Wu et al ., 2019 ). Intriguingly, novel stomatal forms redeploy the core bHLHs in new ways, for example grass stomata are dumbbell-shaped guard cells flanked by subsidiary cells, and MUTE has acquired new roles specifying the identity and position of both guard cells and subsidiary cells ( Raissig et al ., 2017 ; Spiegelhalder et al ., 2024 ; Wu et al ., 2019 ). Stomatal function and development are subject in environmental regulation, and several studies have tracked the evolution of pathways enabling such regulation ( Clark et al ., 2022 ). When environmental response pathways have been described in molecular detail, their impacts on stomatal development typically involve SPCH. In A. thaliana , PHYTOCHROME INTERACTING FACTOR4 (PIF4) represses SPCH expression in warm temperatures ( Lau et al ., 2018 ) and mutations targeting the tomato SPCH promoter render plants unable to change stomatal production in response to light or temperature cues ( Nir et al ., 2023 ). Light and drought affect AtSPCH protein accumulation by modulating the signaling pathways whose downstream kinases phosphorylate AtSPCH ( Kumari et al ., 2014 ; Wang et al ., 2021 ). That AtSPCH is the prime target of environmental regulation may not be surprising given its role as regulator of the asymmetric cell divisions that enable initiation and stem-cell like proliferation of stomatal lineages. Grasses are an economically and ecologically important group of monocot angiosperms. They thrive in dry environments, in part due to their highly efficient stomata featuring dumbbell shaped guard cells and adjacent subsidiary cells ( Franks and Farquhar, 2007 ; Leakey et al ., 2019 ; Linder et al ., 2018 ). Stomata are arranged in linear cell files in the leaf, with each stoma oriented in the same direction. The highly organized progression of stomatal formation begins with an asymmetric cell division, but unlike A. thaliana , there is no stem-cell like phase of repeated divisions, instead the smaller daughter of the asymmetric division is immediately destined to become a pair of guard cells. As in dicots, grass stomatal production can be modified by environmental conditions or precision genetic engineering, the latter of which has been shown to improve water-use-efficiency in a variety of agronomically important cereals ( Caine et al ., 2019 ; Dunn et al ., 2019 ; Hughes et al ., 2017 ; Mohammed et al ., 2019 ). Whether SPCH is the target of environmental tuning in grasses is an open question and is particularly interesting because of (1) the more streamlined development in grass stomatal files and (2) because there are two SPCH paralogs in most grasses. The paralogs BdSPCH1 and BdSPCH2 in B. distachyon and OsSPCH1 and OsSPCH2 in O. sativa are partially redundant in their respective species; loss of either paralog leads to a reduction in stomatal production and loss of both eliminates stomata and leads to seedling death ( Raissig et al ., 2016 ; Wu et al ., 2019 ). Grasses (Poaceae) underwent rapid speciation following the rho whole genome duplication (WGD) ∼140-70 million years ago ( Lovell et al ., 2022 ; Ma et al ., 2021 ; Preston et al ., 2009 ; Wu et al ., 2008 ; Zhang et al ., 2024 ). WGDs have been hypothesized to provide the genetic material for novelty because duplication can release genes from selective pressure and enable them to subfunctionalize or acquire new functions ( Ohno, 1970 ). Broadly, plant WGDs contribute to clade survival and environmental robustness on a macroevolutionary timescale ( Van de Peer et al ., 2009 ). Most plant species return to a diploid state following a WGD ( De Smet et al ., 2013 ; Panchy et al ., 2016 ), yet retain some duplicated genes, and TFs are frequently enriched following plant WGDs ( Almeida-Silva and Van de Peer, 2023 ). Duplicated genes may be retained because they have acquired new functions, they partition ancestral functions, or because gene products are involved in processes where it is critical to maintain stoichiometry ( Birchler and Veitia, 2010 ; Huang et al ., 2022 ; Papp et al ., 2003 ) TF retention may result when paralogous TFs become expressed in different places, acquire new DNA binding site preferences or become incorporated into new gene regulatory networks ( Almeida-Silva and Van de Peer, 2023 ; Gera et al ., 2022 ). The rho WGD provided Poaceae with a complex history of duplicate gene retention and losses, and it is estimated that only about one quarter of duplicated genes were retained ( Zhang et al ., 2024 ). Among the stomatal bHLHs, a second copy of SPCH , but not of MUTE or FAMA , is typically retained in grasses. There are two SCRM paralogs derived from a different duplication event than the Brassica-specific duplication that generated Arabidopsis SCRM and SCRM2 ( Raissig et al ., 2016 ). In this study, we identify the rho WGD as the likely origin of the SPCH duplication and provide evidence that both SPCH paralogs are under selection. Using loss of function mutations and protein reporters for SPCH1 and SPCH2 in the Pooideae member B. distachyon , we show that the two paralogs enable responsiveness to two separate environmental stimuli. To identify the source of paralog-specific regulation, we tested the transcriptional and translational response of each paralog when we altered light and temperature conditions and found that regulation appears to be at the level of protein abundance and persistence in the lineage. Loss of function analyses in the Oryzoideae member, O. sativa are consistent with the SPCH paralogs mediating similar environmental responses across the grasses. Methods Synteny analysis Proteomes, gene locations, and coding domain nucleotide sequences for Hordeum vulgare Morex v3, Triticum aestivum Chinese Spring v2.1, Oryza sativa Nipponbare v7, Zea Mays Ref_Gen v4, Panicum virgatum v5.1, Setaria viridis v4.1, Sorghum bicolor v3.1.1 were downloaded from Phytozome.org (accessed 4/11/2025). Orthofinder v2.5 and MCScanX v1.0.0 were used to identify orthologs and blocks of synteny between the grasses. GENESPACE v1.2.3 was used to visualize the blocks of synteny among grass chromosomes and highlight blocks of synteny around B. distachyon SPEECHLESS ( SPCH ) paralogs BdSPCH1 and bdSPCH2 as a seed for a riparian plot where each species is arranged along the y-axis according to their phylogenetic relationship, accessed from NCBI taxonomy and the regions of synteny. The exclusivity of each of the synteny tracks indicates that the SPCH paralogs duplicated before the speciation of these grasses. Ka and Ks between paralogs To calculate the age of the shared grass SPCH duplication, we used DIAMOND v2.1.11 blastp program to identify similar proteins within each species using the “–more-sensitive” and “--e 1e-10” flags. Results were filtered to top 5 similar genes for each query gene to improve MCScanX results as recommended in the MCScanX methods paper ( Wang et al ., 2012 ). MCScanX identified blocks of collinearity and then calculated Ka (nonsynonymous substitutions per site with the possibility of creating a non-synonymous change) and Ks (synonymous substitutions per site of possible synonymous change) between each pair of paralogs in each species. R (v4.2) with the ggplot2 v2.1 package plotted the density for the Ka and Ks results, colored by species and annotating the values for Ka and Ks for the SPCH1 and SPCH2 paralogs of each species, respectively. Paralog alignment and dN/dS Coding-domain nucleotide sequences for SPCH paralogs from Hordeum vulgare Morex v3, Triticum aestivum Chinese Spring v2.1, Oryza sativa Nipponbare v7, Zea Mays Ref_Gen v4, Panicum virgatum v5.1, Setaria viridis v4.1, and Sorghum bicolor v3.1.1 were downloaded from Phytozome. We used TranslatorX to generate a codon-specific nucleotide multi-sequence alignment across all the paralogs ( Abascal et al ., 2010 ). By creating a nucleotide alignment that preserves codon phase, we could identify the rate of mutations in the nucleotide sequences that lead to synonymous or nonsynonymous changes in the protein. This codon-specific alignment and the phylip gene tree generated by orthofinder were used as inputs into EasyCodeML with the M0 and M2a models( Gao et al ., 2019 ; Yang, 2007). For each paralog, we generated Ka/Ks values across each of these genes and plotted using a custom python script in Matplotlib. A negative score across the length of both paralogs indicates that both are still under purifying (negative) selection. Conservation in cis-regulatory elements (CREs) was explored by downloading 600 bp of sequence upstream of the SPCH1 and SPCH2 TSSs from Phytozome across 9 grass species ( H. vulgare r1, T. aestivum ChineseSpring v2.1, O. sativa Nipponbare v7, P. hallii v3.2, P. virgatum v5.1, S. italica v2.2, S. viridis v4.1, S. bicolor v5.1, Z. mays RefGenV4). Each paralog CRE was aligned with Clustal Omega ( Madeira et al ., 2024 ). We searched for TF motifs in each CRE using FIMO with the JASPAR Core Plant MEME dataset ( Bailey et al ., 2015 ; Rauluseviciute et al ., 2024 ). In addition, we aligned amino acid sequences of SPCH paralogs across the 11 species from assemblies of the 9 grass species above plus A. thaliana (Araport11) and O. Sativa (Kitaake v3.1) using CLUSTALW on https://www.genome.jp/tools-bin/clustalw ( Thompson et al ., 1994 ). We then highlighted the conserved basic helix-loop-helix and SMF (ACT-like) domains in green and yellow, respectively ( Seo et al ., 2022 ). AtSPCH’s previously described PEST domain is indicated in bold letters, highlighted in pink. We also bolded the putative consensus MAPK phosphorylation sites (P-x-S/T-P) in each paralog ( Lewis et al ., 1998 ). Plant Materials Multiple OsSPCH1 and OsSPCH2 knockout alleles were previously generated in Oryza sativa L. japonica cv Zhonghua 11 and described in Wu et al ., 2019 . We used c- osspch1 and t-osspch2 (hereafter osspch1 and osspch2 ) as the representative mutant line for each paralog. CRISPR mutations and a T-DNA insertion were verified by PCR amplification and sequencing (Primers in Table S1). We thank Dr. Suiwen Hou (Lanzhou University) for providing the mutant germplasm. Due to customs constraints and lack of availability of the Zhonghua 11 cultivar in USDA Germplasm Genebank or GRIN-Global database, we were unable to acquire or analyze the Zhonghua 11 wildtype cultivar. We instead analyzed common wildtype cultivars Oryza sativa L. japonica cv Nipponbare and Oryza sativa L. japonica cv Kitaake. We thank Dr. Julia Bailey-Serres (UC Riverside) and Dr. Zhiyong Wang (Carnegie Institution) for seeds. Multiple CRISPR/Cas9 induced knockout alleles of BdSPCH1 and BdSPCH2 were generated previously in B. distachyon line Bd21-3 ( Raissig et al . 2016 ) and we analyzed a single representative line for each paralog. CRISPR mutations were verified through PCR amplification and sequencing (Primers in Table S1). B. distachyon BdSPCH1pro:BdSPCH1-YFP and BdSPCH2pro:BdSPCH2-YFP reporter lines were previously generated in the Bd21-3 line and described in Raissig et al . 2016 . The Bd21-3 line was used as the B. distachyon wildtype. Plant growth Seeds of B. distachyon line Bd21-3, and homozygous bdspch1 , and bdspch2 mutants were sterilized with 1.2% sodium hypochlorite, 0.1% Tween aqueous solution for 15 minutes then rinsed with sterile water before being placed onto sterile 100mm x 15mm square ½-strength Murashige and Skoog (MS, Caisson Labs) agar plates (1% agar, pH 5.7). We stratified the seeds in darkness at 4°C for 4 days before moving them to a regimen of 8 hours light of 50 µE and 16h dark at 22°C in one of two Percival model CU22L growth chambers (serial numbers: 21160.01.15C and 28704.01.20) for another 2 days for germination. After germination, we moved a third of the plates to a higher light treatment of 300uE and a third to a higher temperature treatment of 32°C in a Percival model CU22L growth chamber. Chamber 21160.01.15C was used for testing response to temperature and chamber 28704.01.20 was used for testing response to light intensity. O. sativa seeds were sterilized with 2.62% sodium hypochlorite in an aqueous solution for 30 minutes before being rinsed in water three times. O. sativa seeds were then kept submerged and in darkness at 32°C for 3 days. O. sativa was plated 245 x 25mm square on ½-strength Murashige and Skoog (Caisson Labs MS) agar plates (1% agar, pH 5.7) and moved to Percival model CU22L growth chamber (21160.01.15C) with 8 hour light and 16 hour dark cycles with either 50 µE 26°C, 300 µE 26°C, or 50 µE 32°C conditions. Stomatal phenotyping After the third leaf in B. distachyon and the second leaf in O. sativa fully emerged, we snipped it from the plant and placed it in 7:1 ethanol:acetic acid clearing solution for 1 week. We then mounted leaves on a microscope slide using Hoyer’s solution (7.5 g gum arabic, 5 ml glycerin, 100 g chloral hydrate, 30 ml H2O) and allowed the leaf tissue to clear for one week. DIC images were obtained at 10X for B. distachyon leaves and 20X for O. sativa leaves magnification on a Leica DMi7 microscope, and three regions per leaf were imaged avoiding the midrib in the fully expanded blade. We then measured the leaf area and cell file length with FIJI-ImageJ and counted the abaxial stomata and stomata cell files with the CellCounter plugin ( Schindelin et al ., 2012 ). RT-qPCR B. distachyon ecotype Bd21-3 seeds were germinated on plates as described above with a daily cycle of 8 hours light of 50µE and 16h dark at 22°C in a Percival model CU22L growth chamber. After 16 days, before the emergence of the third leaf, we moved a third of the plates to either 300 µE 22°C or 50 µE 28°C growth chambers. At each time point of 2 hours, 1 day, 2 days, and 3 days after moving them to the new conditions, we dissected the leaf developmental zone by removing the first two leaves and root, then sectioning 4 mm from shoot apical meristem as previously described ( McKown et al ., 2023 ; Raissig et al ., 2016 ). Tissue was harvested from 6 individuals per replicate with 4 replicates per sample then frozen with liquid nitrogen and stored at -80℃ until all samples were collected. We then ground tissue with a Spex Certiprep Geno Grinder 2000 and extracted RNA from each replicate using the Qiagen RNeasy® Mini kit following the manufacturer’s protocol for total RNA extraction from plant cells, including optional DNase treatment. We quantified the RNA using a Nanodrop (ThermoScientific, NanoDrop 1c) and normalized RNA concentrations across samples. We spiked in 10ng of A. thaliana RNA to control for cDNA synthesis variability. To reverse transcribe RNA into single-stranded cDNA, we used the Bio-Rad iScript™ cDNA Synthesis Kit and then quantified the cDNA with Bio-Rad SsoAdvanced™ Universal SYBR® Green Supermix in a CFX96™ Real Time System with primers reported in table S1. Relative expression was computed as 2 −ΔΔ CT by normalizing Ct values to control gene BdUBC18 and reported as relative to the expression of BdSPCH1 or BdSPCH2 in plants remaining at 50 µE 22°C for each the collection time point. Fluorescence microscopy of protein reporters B. distachyon BdSPCH1pro:BdSPCH1-YFP and BdSPCH2pro:BdSPCH2-YFP reporter lines were germinated on plates as described above in a daily cycle of 8 hours light (50 µE) at 22°C and 16h dark in a Percival model CU22L growth chamber. After 16 days, before the emergence of the third leaf, we moved a third of the plates to either 300 µE 22°C or 50 µE 28°C chambers. After 3 days, we carefully removed the third leaf, leaving the developmental zone intact by dissecting it out of the surrounding second leaf. We then stained the cell walls with propidium iodide (1:100 of a 1 mg/mL stock) for 7 minutes in darkness then leaf tissue mounted in water. We then imaged the PI and the YFP at 41X on a Leica Stellaris confocal scanning microscope (513nM excitation with 520-570nM emission bandwidth for YFP and 600-650nM emission bandwidth for propidium iodide). We quantified the YFP expression of SPCH1 and SPCH2 protein reporters within the developing leaf using FIJI-ImageJ to identify and record local positions of YFP intensity maxima along the stomata cell file, drawing a 8µm radius circle, and measuring the integral intensity within the circle ( Schindelin et al ., 2012 ). We measured the length of the zone with fluorescence intensity within each stomata precursor cell file with FIJI-ImageJ. At the base of each stomatal cell file are several cells of uniform shape and size, of which the distal cells will undergo an asymmetric cell division (ACD). The smaller daughter cell of each ACD is fated to become a pair of stomatal guard cells, and at the stages we image leaves, it is possible to see ∼30-50 precursor cells in intermediate stages between the initial ACD and differentiated stoma. We identified the start of the asymmetric cell division (ACD) zone within each cell file as the first YFP+ cell to not be adjacent two other YFP+ cells because ACDs result in larger YFP-cells becoming interspersed in a file. The last YFP+ cell we considered for our measurements was one cell away from the previous YFP+ cell. We ignored the final pre-SCD YFP+ cell in our intensity and length measurements as expression within each cell file typically disappeared several cells before that stage in the lineage before reappearing in that single cell. Results SPEECHLESS paralogs arose from the rho whole genome duplication In previous studies of grass stomatal development, it was noted that SPCH was duplicated, but that MUTE and FAMA, the subfamily Ia bHLHs active at later stages in stomatal development were not ( Raissig et al ., 2016 ; Wu et al ., 2019 ). We were curious whether the two copies of SPCH originated from a small-scale duplication of the chromosome around SPCH , or whether they were retained after WGD while others were lost. The origin of the SPCH duplication would lead us to make different predictions about selective pressures and evolutionary potential of the stomatal lineage gene regulatory networks. Using the regions of B. distachyon chromosomes 1 and 3 that contain BdSPCH1 and BdSPCH2 , respectively as seeds, we could identify blocks of synteny that contain SPCH1 and SPCH2 across the Poaceae family. When visualized in a riparian plot ( Fig. 1a ), SPCH paralogs ( SPCH1 in teal and SPCH2 in red) appear in distinct, non-overlapping block synteny across multiple species suggesting that the paralogs arose from a large-scale duplication that included neighboring genes. Interestingly, H. vulgare (barley) has retained only SPCH1 . Download figure Open in new tab Figure 1: SPCH is retained following the rho whole genome duplication in grasses. (a) Riparian plot showing blocks of synteny containing SPCH1 (teal) and SPCH2 (red) across multiple grass species. Only chromosomes (Chr) with synteny to B. distachyon ’s Chr1 and Chr3 are shown and are scaled by the number of genes. (b) Scaled density plot for Ks between paralogs colored by species. Dashed lines represent the Ks value between SPCH1 and SPCH2 for the species colored. (c) Scaled density plot of Ka. (d) Amino acid alignment of SPCH1 and SPCH2 proteins across multiple grass species. Consensus sequences for each paralog are colored by Ka/Ks ratio for that amino acid. Paralogs from each species are shaded by similarity across paralogs for that amino acid. Green, pink and yellow underlays represent the bHLH, PEST and SMF (ACT-like) domains, respectively, and correspond to the same color-coded regions in the full amino acid alignment in Fig. S1. Based on the synteny analysis and distribution of duplicated SPCH genes we hypothesized that these paralogs arose from the rho WGD. If this were true, then neutral mutation accumulation rates between SPCH1 and SPCH2 should be similar to mutation accumulation rates seen between other genes duplicated in this event. From five grass species with high quality genomes, we identified pairs of paralogs within each genome and then calculated the synonymous mutation rate per synonymous site (Ks) and non-synonymous mutation rate per non-synonymous site (Ka) between each paralog pair ( Fig. 1b,c ). Synonymous mutation rate is often considered to be the rate of neutral evolution ( Ohta, 1992 ). Ks for SPCH paralogs in each species is near the center of Ks distribution for other paralogs in each species, indicating that SPCH duplicated at the same time as the majority of other retained paralogs ( Fig. 1b ). This calculated SPCH Ks (median, 0.68) is consistent with the Ks calculated for Poaceae-wide paralogs that originated in the rho duplication in previous studies ( De La Torre et al ., 2017 ; Tang et al ., 2010 ; Zhang et al ., 2024 ). An exception was observed in Zea mays (maize) where there is an additional paralog of SPCH and the corresponding appearance of a peak at lower Ks values than the other grasses— consistent with the species’ more recent allotetraploid origins ( Fig. 1b , blue) ( Gaut and Doebley, 1997 ). Here, our analysis detected both the pan- Poaceae duplication that created SPCH1 and SPCH2 and a more recent duplication of SPCH2 ( Fig. 1a-c ). The Ks values for the more recent duplication of SPCH2 fall within the peak of Z. mays paralogs ( Fig. 1b ). If the SPCH duplication was a product of a WGD in grasses, then the single copies of MUTE and FAMA were the result of secondary loss, raising the question of why both SPCH copies are retained. Using a metric of selection, the Ka to Ks ratio, we found that non-synonymous mutations are less frequent than synonymous mutations in SPCH1 and SPCH2 , indicative of purifying (negative) selection on both genes ( Fig. 1c ). To identify sites in each gene likely to be the targets of purifying selection, we examined mutation rates separately across SPCH1 and SPCH2 paralogs among several Poaceae species, and identified several regions that are conserved and under purifying selection ( Fig. 1d ). The most conserved of these align with domains shown to be important for function in the single SPCH of A. thaliana including the bHLH domain that enables sequence-specific DNA binding ( Davies and Bergmann, 2014 ; Lau et al ., 2014 ) and a C-terminal ACT-like domain essential for SPCH activity ( Davies and Bergmann, 2014 ; MacAlister and Bergmann, 2011 ) that can mediate protein-protein interactions ( Figs 1d ,S1) ( Seo et al ., 2022 ). In addition, alignment of the proximal 5’ regulatory regions of SPCH paralogs across grass species show that SPCH1 and SPCH2 diverged considerably, but that there are blocks of conservation immediately upstream of the TSS of each that contain binding sites of known developmental regulators (Fig. S2). Different environmental responsiveness exhibited by B. distachyon ’s SPCH paralogs SPCH1 and SPCH2 acquired molecular differences in both the coding region and within the cis-regulatory regions since their duplication. Because these differences between the paralogs were conserved after speciation across many grasses, we hypothesized that the paralogs acquired functional differences that kept each paralog important and unique. Evidence that both paralogs are under selection is interesting in light of functional studies in B. distachyon and O . sativa showing that SPCH1 and SPCH2 are expressed in and involved in stomatal development, but show unequal redundancy–that is SPCH2 has a quantitatively larger effect on stomatal production than SPCH1 and loss of both eliminates stomata completely ( Raissig et al ., 2016 ; Wu et al ., 2019 ). SPCH has a key role integrating environmental conditions into tuning the stomatal lineage in the dicot A. thaliana , and so we hypothesized that grass SPCH paralogs could sub-functionalize their roles in environmental sensing. To test the ability of SPCH1 or SPCH2 to mediate a change in stomatal production in response to environmental perturbation, we needed to be able to assay plants with only one or the other functional paralog and thus turned to single gene disruption alleles (knockouts) previously characterized in B. distachyon ( Raissig et al ., 2016 ). If either gene mediated the stomatal density (SD, stomata/unit area) response to environmental perturbation, we expected the response to be abrogated in that mutant. In grasses, stomata develop in specific epidermal cell files, and stomata density is a product of the number of cell files in which stomata form and the number of stomata in a given cell file. To measure SD we harvested fully expanded 3rd leaves and imaged the epidermis along the leaf blade, focusing on the developmental zone near the base and avoiding the midrib. For each region imaged, we quantified the number of stomata, the number of stomatal cell files, and the length of each cell file from the initiation to termination in our micrographs to estimate the average distance between stomata in a single cell file ( Fig. 2a ). Download figure Open in new tab Figure 2: Plants bearing mutations that leave only one SPCH paralog functional cannot adjust stomatal density to environmental changes. (a) DIC images of B. distachyon abaxial leaf epidermis. Stomata are false-colored green. Images are arranged in rows by environmental conditions and columns by genotype. Scale bar is 50μm. (b) Stomatal density on the abaxial leaf epidermis for plants grown under 22°C or 28°C, 50µE conditions. Plants lacking BdSPCH1 ( spch1 ) are not sensitive to the temperature change. (c) Stomatal density on the abaxial leaf epidermis for plants grown under 50µE or 300µE, 22°C conditions. Plants lacking BdSPCH2 ( spch2 ) are not sensitive to the light intensity change. (d) Average distance (interval) between two stomata in a row. Stomatal spacing decreases with light intensity except in spch2 plants. (e) The number of epidermal cell files that produce stomata (row density) increases with increased temperature except in spch1 plants. In (b-e) each point represents a measured developing leaf region (n > 15 regions per genotype and condition). Boxes in panels (b-e) indicate the median (horizontal line) and the interquartile range (box: 25th–75th percentiles). Significance measured by two-way ANOVA with Tukey post-hoc reporting the effect of condition while controlling for leaf identity of each region (* < 0.05, **** < 1e-04). We established the wildtype Bd21-3 stomatal density response to growth under three different light and temperature regimes (22°C 50 µE, 22°C 300 µE and 28°C 50 µE). The growth conditions generated by changing light intensity and temperature had reproducible effects on stomatal development, with wildtype increasing stomatal density in response to increasing light (50 µE to 300 µE) or temperature (22°C to 28°C) ( Fig. 2a-c ). In contrast, bdspch1 mutants failed to show a significant increase in stomatal density in response to warm (28°C) temperature ( Fig. 2b ) but still responded to an increase in light intensity ( Fig. 2c ). Conversely, bdspch2 mutants increased their stomatal density in response to a temperature increase ( Fig. 2b ) but were insensitive to increased light intensity ( Fig. 2c ). These reciprocal insensitivities suggest that B. distachyon SPCH paralogs have acquired paralog-specific environmental responsiveness. The different insensitivities also allowed us to eliminate the concern that the overall lower number of stomatal in each of the single mutants would render environmental response difficult to generate or to measure. Even in bdspch2 where overall stomatal production is substantially lower, a response to a change in temperature can be detected. We found that the cellular mechanism by which SD increased differed between light and temperature change treatments. The distance between stomata in a single cell file decreased significantly when we increased light intensity, but not when we increased temperature ( Figs 2d , S3a), except in bdspch2 where the reverse is true. An increase in the density of stomatal-forming files drove the increase in response to increased temperature (except in insensitive bdspch1 ) but did not change when we increased light intensity ( Figs 2e ,S3b). Again, the differential behaviors of bdspch1 and bdspch2 under these conditions can help pinpoint where in the pathway–from perception of environment to developmental response–the paralogs differ. For example, although bdspch2 does not decrease spacing in a row in response to a change in light intensity, it can decrease spacing in response to alteration in temperature. Thus, it is likely that SPCH1 and SPCH2 can participate in driving similar downstream developmental pathways, and that there is difference in how the upstream environmental cues regulate the paralogs. Expression and protein stability of B. distachyon paralogs in response to environmental conditions We then investigated two ways that upstream environmental cues could differentially regulate BdSPCH1 and BdSPCH2 . The first hypothesis was BdSPCH1 and BdSPCH2 would be transcribed under different conditions, an idea in line with the literature that has shown rapid adaptation through cis-regulatory differences ( Fraser et al ., 2010 ; Mack et al ., 2018 ). Our alignment of 5’ regulatory sequences supported a model that BdSPCH1 and BdSPCH2 could be targets of different TFs (Fig. S2), and cell-type specific RNA-seq experiments show that BdSPCH1 and BdSPCH2 levels differ across organs and life stages ( Hao et al ., 2021 ). In Arabidopsis, a stomatal density temperature response is mediated through repression of SPCH transcription by the thermosensory TF PIF4 ( Lau et al ., 2018 ). To test for a SPCH transcriptional response to our alterations of temperature or light, we grew wildtype B. distachyon under our standard temperature and light conditions, then moved seedlings to higher temperature or higher light intensity for increasing lengths of time (2 hours to 3 days). BdSPCH1 and BdSPCH2 are only expressed in the developing leaf zone ( Raissig et al ., 2016 ) so we extracted RNA from dissected tissue from this zone. RT-qPCR-based expression profiles showed that RNA abundance of both BdSPCH paralogs changed in the same way in response to manipulations of light intensity or temperature at each of the time points we collected following transfer ( Fig. 3a ). Therefore, we find no evidence that the difference stomata density responses to environmental change observed in the bdspch1 vs. bdspch2 is a consequence of different transcriptional regulation of the paralogs. Download figure Open in new tab Figure 3: Differential environmental response appears to affect the persistence of SPCH protein, not level of SPCH expression. (a) RT-qPCR quantification of BdSPCH1 and BdSPCH2 RNA expression after moving B. distachyon Bd21-3 plants acclimated to 50µE and 22°C to 300µE light or 28°C temperature for the times specified on the X-axis. Expression quantification was normalized to a control gene and then to the expression of SPCH1 or SPCH2 isolated from unshifted plants. Bars represent Standard Error (SE). (b) Cartoon and confocal images illustrating B. distachyon stomatal lineage development. For this study, BdSPCH2:BdSPCH2-YFP nuclear accumulation (yellow) is quantified from the first asymmetric division (bottom inset, “first ACD”) until the final cell in the contiguous expression domain (top inset, “last measured”). In confocal images, cell outlines are in magenta. Scale bar is 20μm. The cartoon truncates the length of the full developmental zone and ignores hair cells for simplicity. (c,d) Length of detectable BdSPCH1:BdSPCH1-YFP or BdSPCH2:BdSPCH2-YFP fluorescence in “measured zone”. Dots represent cell files across multiple leaves (n > 8 per reporter per condition). Boxes in panels (c) and (d) indicate the median (horizontal line) and the interquartile range (box: 25th–75th percentiles). Significance measured by two-way ANOVA with Tukey post-hoc reporting the effect of condition while controlling for leaf identity of each cell file (** < 0.01, *** < 1e-3, **** < 1e-4). We therefore tested our second hypothesis, that paralogs are subject to post-translational regulation. Again, there is precedence for this mechanism in the response of Arabidopsis SPCH to environmental factors ( Kumari et al ., 2014 ; Wang et al ., 2021 ) and the grass SPCH1 and SPCH2 alignments show differences in the presence and position of potential phosphorylation sites ( Figs 1d , S1) ( Raissig et al ., 2016 ). We imaged previously generated BdSPCH1pro:SPCH1-YFP and BdSPCH2pro:SPCH2-YFP reporters, and consistent with the prior publications generating these materials ( Raissig et al ., 2016 ), we detected expression of SPCH1-YFP and SPCH2-YFP in stomatal precursors ( Figs 3b ,S4a-d). SPCH2-YFP expression appeared in early proliferative cell files and became much stronger in the nuclei of the smaller daughter cells resulting from asymmetric cell divisions (ACDs) in the stomatal cell files. SPCH1-YFP was only detectable after the onset of ACDs. Both SPCH1-YFP and SPCH2-YFP expression decreased as cells matured, but we also noticed a transient burst of nuclear YFP just before the symmetric cell division that creates guard cells (arrow in Fig. S4c,d). This pulse of expression provided a fiducial mark to measure stomatal cell files, but we did not include this cell in our quantifications because fluorescence in the lineage typically disappears a few cells before then (Fig. S4c,d). Under the same experimental regime as used for assaying transcriptional response, we did identify paralog-specific protein reporter responses to the changes in light intensity and temperature. Specifically, SPCH1-YFP fluorescence persisted longer in stomatal cell files in plants moved to a higher temperature (22°C to 28°C) for 3 days compared to plants moved to a higher light intensity (50 µE to 300 µE) or that remained in the original conditions ( Fig. 3c ). SPCH2-YFP fluorescence persisted the longest in plants moved to higher light conditions ( Fig. 3d ). SPCH2-YFP fluorescence could also be detected in proliferating cells at the leaf base but we restricted our quantification to after the first ACD ( Fig. 3b ) to compare to SPCH1 and focus on confirmed stomatal cell behaviors. What does extended persistence of BdSPCH reporters mean? In the context of a stomatal cell file, this could be an indication of a prolonged meristematic capability. It could also be a consequence of higher protein accumulation in each cell. By measuring reporter fluorescence intensity per cell, however, we found that the persistence of SPCH1-YFP and SPCH2-YFP was not just due to higher per cell intensity (Fig. S5a,b). In addition, when tracking SPCH2-YFP intensity along a stomatal precursor cell file from base to tip, we found that, under all tested environmental conditions, intensity followed the same low, high, low pattern, with intensity increasing in the middle of the file before trailing off as cells matured (Fig. S5c). We tested whether the length of persistence within the cell file was a consequence of expanding the distance between each stomata precursor. Spacing between the last 10 SPCH1-YFP cells did increase under high temperature conditions, but spacing of the last 10 SPCH2-YFP cells did not significantly increase in any of the environmental perturbations (Fig. S4a,b). Increasing the length of persistence decreases the number of cells between the final pre-SCD cell and second-to-last cell with expression within the cell file (Fig. S4c, distance between yellow bracket and yellow arrowhead). Overall, reporter expression persistence in stomatal cell files matches the expectations from loss of function mutations as to which SPCH paralog mediates the stomatal density response to our increased light or temperature treatments (e.g., SPCH2 shows longer persistence in higher light and spch2 mutants were insensitive to increased light). This suggests that environmental regulation specifies not how strongly SPCH paralogs are expressed, but the ability of their proteins to be maintained through development. In Arabidopsis, SPCH protein stability is modulated by MAPK target sites and/or a PEST domain ( Davies and Bergmann, 2014 ; Lampard et al ., 2008 ); paralog-specific differences in the presence of these sequences in SPCH1 and SPCH2 ( Figs 1d , S1) indicate the potential for this regulatory mechanism to differentially affect paralogs in the grasses. Paralog-specific responses to light and temperature are also seen in O. sativa SPCH duplicated before the speciation of B. distachyon so we were curious whether this subfunctionalization of environmental sensitivity in stomata development was conserved in other grasses that retained both paralogs following the rho WGD. O. sativa (rice) and B. distachyon diverged ∼101-53 MYA and belong to different tribes within the same BOP clade (as distinct from PACMAD containing Zea mays and Sorghum bicolor ) ( Christin et al ., 2014 ; Zhang et al ., 2024 ). Mutations that eliminate OsSPCH1 or OsSPCH2 function were previously generated in O. sativa ( Wu et al ., 2019 ) and genomes of several O. sativa cultivars with sequence variation around SPCH loci are available as well ( Fujino et al ., 2015 ; Jain et al ., 2019 ; Matsumoto et al ., 2016 ), thus we tested stomatal development in response to environmental perturbation in loss of function osspch1 and osspch2 mutants, and in the O. sativa L. japonica cultivars Nipponbare and Kitaake. In O. sativa , loss of OsSPCH2 results in a severe reduction in stomatal density, but loss of OsSPCH1 was reported to have no discernible stomata density defect ( Wu et al ., 2019 ). We replicated these results in our chambers and standard growth conditions ( Fig. 4a,d,e ). We then challenged the OsSPCH mutants and Nipponbare and Kitaake cultivars to light and temperature shifts paralleling the shifts performed on B. distachyon , but using the reported optimal O. sativa growth temperature of 27°C as baseline and 32°C as warm temperature. We compared the responsiveness of osspch1 and osspch2 mutants and found that osspch2 was insensitive to increasing light intensity. Nipponbare, Kitaake and osspch1 responded by decreasing stomatal density ( Fig. 4a-e ). The stomata density response to growth in higher light conditions is opposite in direction from that in B. distachyon plants, but both osspch2 and bdspch2 are insensitive. In response to increased temperature, osspch2 but not osspch1 was able to increase stomatal density ( Fig. 4a,d,e ). Thus, SPCH1 and SPCH2 seem to retain similar environmental responsiveness among different species within the BOP grass subfamily. Comparing the behaviors of Kitaake and Nipponbare cultivars in the temperature shift experiment, however, introduced other potential explanations for environmental insensitivity; namely that Kitaake responded similarly to osspch1 in not responding to increased temperature, leaving the possibility of a cultivar-specific sensitivity to temperature increase in developing leaves ( Fig. 4b,c ). OsSPCH1 sequences from Kitaake and Nipponbare differ in the presence of a high stringency MAPK target site, providing a potential explanation for the effect, but future experiments would be required to test this (Fig. S1). Download figure Open in new tab Figure 4: Loss of function OsSPCH mutants exhibit the same environmental insensitivities seen with mutations in B. distachyon SPCH paralogs (a) DIC images of O. sativa abaxial leaf epidermis. Stomata are false colored green. Images are arranged in columns by environmental conditions and rows by genotype. Scale bar is 50μm. (b-e) Stomatal density on the abaxial leaf epidermis for plants grown under different light or temperature regimes. OsSPCH1 knockout (KO) and Kitaake plants are not sensitive to the temperature change and OsSPCH2 KO plants are not sensitive to the light intensity change. Dots represent regions across multiple leaves (Kitaake n > 7, others n > 16). Boxes in panels (b-e) indicate the median (horizontal line) and the interquartile range (box: 25th–75th percentiles). Significance measured by two-way ANOVA with Tukey post-hoc reporting the effect of condition while controlling for leaf identity of each region (* < 0.05,** < 0.01, *** < 1e-3, **** < 1e-4). Interestingly, when we tracked the cellular mechanism underlying stomatal density adjustments, we found that in O. sativa genotypes that respond to changing light intensity, it was due to changing the number of stomatal rows instead of the spacing between stomata within a row (Fig. S6a,b). This differs from the cellular mechanism of response in B. distachyon ( Figs 2d ,S3b). In response to temperature, all O. sativa genotypes except Kitaake decrease the distance between stomata within a cell file (Fig. S6a), also different from the observed B. distachyon behavior ( Fig. 2e ,S3a). Discussion The potential for plants to regulate both stomatal aperture and stomatal development enables them to optimize their water use efficiency and photosynthetic capacity across many environments. We show for two representative grasses that stomatal density appears to be tuned in response to environmental cues through varying the number of cell files capable of producing stomata and the number of stomata per file. Changes in stomatal production correlate with differential persistence of SPCH protein accumulation in stomatal files, with SPCH1 and SPCH2 exhibiting unique responses to different environmental conditions. Moreover, we show retention of both copies of the SPCH TF originating in the rho WGD predating Poaceae speciation may mediate subsets of this response in a conserved manner across BOP clade grasses. Several mechanisms have been proposed to support the retention of paralogs following WGD in plants including neofunctionalization, subfunctionalization, dosage balance, or neutral variation ( Birchler and Yang, 2022 ; Freeling, 2009 ; Iohannes and Jackson, 2023 ). These models can interact and complement each other, for example, dosage balance can maintain redundant paralogs until they acquire subfunctionalization through neutral evolution ( Iohannes and Jackson, 2023 ). While many paralogous TFs were retained from the rho WGD, SPCH is unique among stomatal lineage bHLH TFs in both copies being retained in most grasses. Each SPCH paralog exhibits molecular signatures of purifying selection in the protein coding region ( Fig. 1b,d ) suggesting an active mechanism maintaining paralogs ( Zhang et al ., 2024 ). Are any of the proposed general models for paralog retention likely to apply to SPCH ? A dosage balance model where both SPCH paralogs are maintained because reduction in expression would be maladaptive does not fit with previously reported expression data that shows the paralogs have different ratios of expression at different developmental stages (e.g., in embryo vs. along the developmental zone in leaves) ( Hao et al ., 2021 ; Zhang et al ., 2022). An extension of this model would suggest that the paralogs would be retained to maintain stoichiometric balance between paralogs and their binding partners ( Birchler and Yang, 2022 ; Kuzmin et al ., 2020 ), but this also seems insufficient because the heterodimer partners of SPCH paralogs, SCRM1 and SCRM2, share a more ancestral duplication with other Poales like Ananas comosus and are not thought to originate in the rho duplication. The phenotypic effects of loss of function mutations in either SPCH1 or SPCH2 affect stomatal density rather than other processes, consistent with subfunctionalization rather than neofunctionalization contributing to the retention of these paralogs since the rho WGD ( Raissig et al ., 2016 ; Wu et al ., 2019 ). Functional tests in B. distachyon BdSPCH1 and BdSPCH2 mutants grown in different environments support the model of subfunctionalization. Specifically, the paralogs differ in their contributions to the stomatal development response to increasing light intensity or temperature, with SPCH1 enabling response to the increased light treatment and SPCH2 to increased temperature. Either paralog could engage the same downstream changes in the density of stomatal cell files or distance between stomata in a cell file, suggesting that it is more likely that the paralogs are subject to different environment-mediated regulation than that they have become enmeshed in different gene regulatory networks to regulate downstream developmental programs. While we demonstrate subfunctionalization between BdSPCH paralogs, we understand a limitation of our approach is that we cannot discern the causal environmental factor in the paralog-specific response to temperature or light intensity. Increasing temperature will increase water vapor pressure and likely increase humidity—additional abiotic factors that have known stomatal density responses ( Tricker et al ., 2012 ; Devi & Reddy, 2018 ). In addition, increasing light intensity and temperature both impact photosynthesis within the leaf by either modulating biochemical rates or deactivating the pathway at more extreme conditions ( Zhang et al ., 2002 ; Scafaro et al ., 2023 ). Subsequently, available sugar influences stomata development and decreases stomata density (for example, Gong et al ., 2021 ). Regardless of the identity of environmental covariates perturbed by light intensity or temperature changes, plants with mutations inactivating SPCH1 and those with mutations inactivating SPCH2 have distinct stomatal responses to the two perturbations, consistent with the SPCH paralogs undergoing subfunctionalization. Several studies of plant homologs have focused on a transcriptional basis for subfunctionalization following a WGD, thereby expanding our understanding of the evolutionary consequences of duplication ( Harikrishnan et al ., 2015 ; Li et al ., 2025 ). These studies build upon the classic literature tying rapid evolution of regulatory genes to adaptive mutations in cis-regulatory regions ( Fraser et al ., 2010 ; Mack et al ., 2018 ). Indeed, we identified conserved cis-regulatory regions that differ between SPCH paralogs across the grasses containing distinct TF motifs (Fig. S2) that could suggest a transcriptional mechanism to explain how paralogs differently affect the phenotype. However, our measured transcriptional differences were insufficient to explain the different dependencies of light and temperature response on the SPCH paralogs in B. distachyon ( Fig. 3a ). Neither were average translational intensities sufficient to explain the paralog-specific response to the environment (Fig. S5a,b). Instead, we identified changes in the zone of protein expression that matched the phenotypic behaviors we observed; the zone of protein accumulation within a stomatal file uniquely increased in the reporter of the paralog implicated in mediating a certain environmental response when the plants were exposed to that condition ( Fig. 3c,d ). Testing if this subfunctionalization was shared across the BOP clade with O. sativa mutants and cultivars revealed a conserved paralog-specific role for SPCH1 and SPCH2 in response to temperature and light intensity perturbations, respectively. We note insensitivity to temperature increase seen in osspch1 was shared with one of two wildtype cultivars we tested–allowing for the possibility that the temperature insensitivity originates in the background cultivar from which osspch1 was isolated. This explanation, however, would require loss of OsSPCH2 to have a gain-of-function effect, newly acquiring temperature sensitivity. In A. thaliana , different ecotypes display different stomata development responses to changing temperature ( Nir et al ., 2023 ). The cultivar-specific response underscores the need to test multiple species/cultivars when exploring the environmental regulation of developmental pathways. Here we present the biological phenomenon of stomatal development responses to environmental change and how it may be mediated through the duplicated SPCH genes. The details of how the paralogs can be regulated and how they differ in their regulation is an opportunity for future studies. For example, could this subfunctionalization be adaptive in increasing the species’ environmental tolerance? One approach is to look for correlations between the copy number of SPCH paralogs and thermal or latitude ranges in grasses that retained the duplication and the few grasses that did not. This approach of correlating SPCH duplications with environmental species range could be expanded across angiosperms–exploring the impact of additional independent duplications of SPCH ( Danzer et al ., 2015 ; McKown et al ., 2019 ; Menezes et al ., 2025 ) and contributing to our understanding of why TFs involved in environmental responses have increased retention rate following WGD in plants ( Panchy et al ., 2019 ; Rody et al ., 2017 ). For example, Populus trichocarpa species have two copies of SPCH derived from an independent (from rho ) duplication, and it appears that alleles of one paralog now predict the ratio of stomata present on the adaxial and abaxial leaf surfaces along an environmental gradient ( McKown et al ., 2019 ). Similarly, our study inspires us to consider further functional studies in members of Poaceae that inherited the SPCH duplication but secondarily lost a paralog. Hordeum vulgare (barley) and Triticum aestivum (wheat) are close relatives of B. distachyon in the same subfamily, the “core” Pooideae, but both lost their SPCH2 paralog and hexaploid wheat now has three copies of SPCH1 . Are barley and wheat stomatal densities insensitive light intensity changes? If they are light sensitive, could their SPCH1 paralog have re-acquired post-translational regulation observed in Bd SPCH2 or OsSPCH2 or have they adopted novel modes of transcriptional regulation? Finally, the ability to assay reporters of BdSPCH1 and BdSPCH2 protein behavior was essential to consider molecular mechanisms underlying paralog differences. Introducing functional protein reporters of grass SPCH1 and SPCH2 into A. thaliana spch mutants could reveal whether the grass paralogs still differ in their capacity to mediate light and temperature stomatal density responses. This heterologous system could also facilitate more detailed investigation of the specific protein domains and residues that mediate environmentally-tuned behavior, with MAPK target sites and a protein-stability determining PEST domain ( Lampard et al ., 2008 ; Raissig et al ., 2017 ; Rogers et al ., 1986 ) being strong candidates to test due to their pattern of being conserved among orthologs and distinct between paralogs across grass species (Fig. S1). Our study contributes a careful examination of the role that paralogs can play in environmental acclimation through non-transcriptional regulation and identifies a novel subfunctionalization in grass paralogs of SPCH , a major stomatal development regulator. This contributes to a growing body of knowledge about the mechanisms by which agronomically important plants have or could modify stomatal development in response to changing climates and may contribute to the development of solutions to the challenge of ever-increasing environmental stressors affecting our crop systems. Author contributions JME contributed to conceptualization, data acquisition, analysis and interpretation and wrote the manuscript in collaboration with DCB. BB contributed to data acquisition. DCB supervised the work. Competing interests The authors declare no competing interests Data availability The data that support the findings of this study, including proteomes, gene locations, and coding domain nucleotide sequences were obtained from Phytozome.org and details of SPCH homologues are provided in Table S2. No new sequence data were generated. Acknowledgements We thank Zhiyong Wang, Julia Bailey-Serres and Suiwen Hou for rice seeds and Alex Borowsky for advice about rice cultivation. Anastacia Del Rio helped with initial characterization of environmental response and Charlie Hale (Cornell) advised on identifying TF motifs in CRE. Jana Sipkova advised on image processing in ImageJ-FIJI. We thank lab members Katelyn McKown and Gabe Amador for discussions and Alysse Pusey and Genevieve Stier for plant care. JME was supported by funds from the National Institutes of Health (T32GM007276). DCB is an investigator of the Howard Hughes Medical Institute. Funder Information Declared National Institutes of Health , T32GM007276 Howard Hughes Medical Institute, https://ror.org/006w34k90 Footnotes Minor modifications to text and figures to improve clarity. References ↵ Abascal F , Zardoya R , Telford MJ . 2010 . TranslatorX: multiple alignment of nucleotide sequences guided by amino acid translations . 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