Nucleotide sequences for gene synthesis were designed from the predicted cDNA sequences of ChECs that were previously identified from ESTs and the fungal genome sequence (Kleemann et al., 2012; O’Connell et al., 2012). Selected ChEC genes were synthesized using GeneART^® gene synthesis technology without their predicted fungal signal peptides and cloned into the pDONR221 GATEWAY entry vector (Invitrogen, Darmstadt, Germany). The sequences were then transferred using the LR cloning reaction into binary destination vectors derived from pSITE (Martin et al., 2009) for transient expression of the proteins in plants as N-terminal fusions with EGFP (pSITE-2CA) or mRFP (pSITEII-6C1). After the LR reaction, the constructs were cloned into Escherichia coli TOP10 cells and then Agrobacterium tumefaciens (C58C1, pGV2260, or pMP90). E. coli cells were cultivated at 37°C in lysogenic broth (LB) medium (Bertani, 1951). A. tumefaciens strains were cultivated on LB at 28°C, with appropriate antibiotic selection (spectinomycin 100 μg ml^-1, streptomycin 50 μg ml^-1, rifampicin 50 μg ml^-1).

The expression patterns of the selected ChEC genes were profiled using RNA-Seq data corresponding to four developmental stages of C. higginsianum, namely in vitro appressoria (22 h post inoculation, hpi), in planta appressoria (22 hpi), biotrophic phase (40 hpi), and necrotrophic phase (60 hpi). Preparation of the RNA samples was described previously (O’Connell et al., 2012) and the raw data sets are available under GEO accession GSE33683. Here, the filtered reads were mapped onto the reannotated genome assembly of C. higginsianum (Dallery et al., 2017) using TopHat2 (Kim et al., 2013) (version 2.0.14, I = 5000, a = 10, g = 5). Previously described ChECs (Kleemann et al., 2012) that lack a gene model in the new annotation were manually incorporated into the annotation file used for mapping. HTseq (Anders et al., 2015) (version 0.5.3p9) was used to count the mapped reads, and ‘Relative Expression Index’ and gene expression level were calculated as described previously (Hacquard et al., 2016). Heatmaps were produced using Genesis software, version 1.7.6 (Sturn et al., 2002).

Nicotiana benthamiana plants were grown under long-day conditions (16 h light, 25°C, 80% relative humidity; 8 h dark, 22°C, 50% relative humidity). An over-night culture of A. tumefaciens strain C58C1 pGV2260 was centrifuged, resuspended in buffer (10 mM MgCl₂, 10 mM MES-KOH pH 5.7, 150 μM acetosyringone) and incubated for 3 h at room temperature. The bacterial suspension was adjusted to OD_{600 nm} = 0.4 and pressure-infiltrated into the abaxial surface of N. benthamiana leaves of 4-week-old plants using a needleless syringe (1 ml). For co-expression experiments, Agrobacteria harboring each construct were mixed and adjusted to OD_{600 nm} = 0.4 in the same suspension and infiltrated. To ensure high levels of transient expression of both constructs in these experiments, we also co-expressed the P19 gene, a suppressor of post-transcriptional gene silencing from the Tomato bushy stunt virus (Oh et al., 2009). Agrobacteria harboring P19 were infiltrated at a final OD_{600 nm} of 0.1. At 24–48 h after agro-infiltration, pieces of leaf tissue (∼5 mm × 5 mm) were excised for examination by confocal microscopy (described below).

The protocol was adapted from that of Marion et al. (2008). Briefly, A. thaliana seeds were surface-sterilized by washing in ethanol (70% v/v) for 1 min and sodium hypochlorite (commercial bleach, 3% v/v) for 15 min and then rinsed five times in sterile water. Seeds were then placed into the wells of a sterile 6-well plate, each well containing 4 ml of half-strength Murashige and Skoog medium covered with a sterile nylon mesh disk (200 μm mesh size). Plates were maintained at 4°C overnight before being transferred to a controlled environment chamber (12 h photoperiod, 230 μmole m^-2 s^-1 photon flux density, 23°C). Transformation was performed on 4-day-old seedlings. An over-night culture of A. tumefaciens strain C58C1 pMP90 was centrifuged and resuspended in buffer (10 mM MgCl₂, 10 mM MES-KOH pH 5.7, 150 μM acetosyringone) and incubated for 3 h at room temperature. Seedlings were completely immersed in the bacterial suspension and then vacuum-infiltrated twice for 1 min. The bacterial suspension was then removed by pipetting and the plants transferred to a growth chamber for a further 3 days. Cotyledons were excised prior to mounting in water under a coverslip for confocal microscopy.

To determine the subcellular localization of the GFP-tagged ChECs in N. benthamiana leaf pieces or A. thaliana seedlings, samples were first mounted under a coverslip in water inside glass-bottomed culture dishes (40 mm diameter, 0.17 mm glass thickness, WillCo-Dish^®) and then observed using either Leica SPE or Leica SP5 inverted confocal microscopes equipped with a ×63 (1.2 NA) water immersion objective. To image GFP fluorescence, the 488 nm laser line was used for excitation and emission was collected between 490 and 525 nm. For imaging mRFP fluorescence, excitation was at 532 nm and emission was observed between 580 and 650 nm. To minimize spectral bleed-through between fluorescence channels during co-localization experiments, images were acquired by sequential scanning with alternation between frames (Leica SPE microscope) or between lines (Leica SP5 microscope). Plant nuclei were stained using DAPI (4’,6-diamidino-2-phenylindole, 10 μg/ml in water) pressure-infiltrated into the abaxial side of the leaf using a 1 ml syringe 30 min before observation. DAPI fluorescence was excited at 405 nm and observed between 440 and 475 nm. Measurement of nuclear areas was performed on maximum projections of confocal image stacks using the Fiji particle analysis tool after manual thresholding (Schindelin et al., 2012). Incomplete nuclei located at image boundaries were manually excluded from these analyses.

Pieces (2 mm × 3 mm) of N. benthamiana leaves (2 days after agro-infiltration) were fixed for 2 h in a mixture of 4% (w/v) para-formaldehyde and 0.5% (v/v) glutaraldehyde in 0.05 M sodium cacodylate buffer (pH 6.9). After dehydration in a graded water–ethanol series, the samples were embedded in LR White acrylic resin using the progressive lowering of temperature method (Satiat-Jeunemaitre and Hawes, 1992). Adjacent ultrathin sections were mounted on separate specimen grids for labeling with different antibodies. Immunogold labeling was as described by Kleemann et al. (2012) except that GFP was detected using a rabbit polyclonal anti-GFP antibody (ab6556, Abcam, diluted 1:50) and DsRed was detected using a rabbit polyclonal anti-RFP antibody (R10367, Molecular Probes, diluted 1:250). Goat anti-rabbit antibodies conjugated with 10 nm colloidal gold particles (British Biocell International, Cardiff, United Kingdom) were used as secondary antibodies. Sections were stained with 2% uranyl acetate for 10 min followed by lead citrate for 15 min and examined with an Hitachi H-7650 TEM operating at 100 kV fitted with an AMT XR41 M digital camera.

Agro-infiltrated N. benthamiana leaves (2 days after infiltration) were first checked for expression of GFP-ChEC fusion proteins by epi-fluorescence microscopy and then snap-frozen in liquid nitrogen. The frozen leaves were reduced to powder using a cold mortar and pestle. Protein extracts were prepared as described by Petre et al. (2015) and 15 μl samples were run on 12% SDS-PAGE gels. Protein concentrations were estimated by Ponceau staining. Proteins separated by SDS-PAGE were transferred to polyvinylidene difluoride membranes (Bio-Rad #1620177) using a Trans-blot Semi Dry transfer cell (Bio-Rad) in transfer buffer (Tris 25 mM, glycine 192 mM, ethanol 20% v/v, SDS 0.1% w/v, pH 8.3). After blocking in 3% w/v bovine serum albumin (BSA) in TBST (Tris-buffered saline containing 0.1% v/v Tween 20), the membranes were probed with mouse anti-GFP monoclonal antibodies (Santa Cruz F56-6A1) diluted 1:5,000 in 3% BSA for 1 h, followed by goat anti-mouse polyclonal antibodies conjugated with horse radish peroxidase (HRP, Dako P0447) diluted 1:20,000 in 3% BSA for 1 h. Blots were developed using Immobilon Western HRP substrate (Millipore) for chemiluminescence detection.

We previously identified genes encoding potential effector proteins from C. higginsianum using two independent approaches. Based on the fungal genome annotation, we identified 365 Candidate Secreted Effector Proteins (CSEPs) defined as extracellular proteins (WoLF PSORT prediction) without homology to proteins from organisms outside the genus Colletotrichum (O’Connell et al., 2012). Based on mining EST data from deep-sequencing in planta cDNA libraries, we found 102 ChECs based on the presence of an N-terminal signal peptide (SignalP prediction) and their lack of homology to known proteins, or similarity to effectors from other fungi (Kleemann et al., 2012) (Figure 1A). In the present study, data from both approaches were combined to obtain a non-redundant list of effectors, hereafter referred to as ChECs. We selected candidates that were preferentially expressed in planta by appressoria and/or biotrophic hyphae by filtering available RNA-Seq data (O’Connell et al., 2012) to include only genes with <15% of total reads derived from a necrotrophic phase library and >50% reads derived from penetrating appressoria and/or biotrophic phase libraries. Genes with low expression levels were discarded if the total number of mapped reads across all three in planta infection stages was <50. Finally, all gene models were manually curated by comparison to the transcriptome data to correct errors in start/stop sites, intron structure and sequence indels. Following this procedure, a total of 61 ChECs were selected for further study (Figure 1A and Supplementary Table 1).

To search for the presence of homologous proteins in other fungi, we blasted the protein sequences of the 61 ChECs against the UniProt database (Swiss-Prot + trEMBL, version 02/03/2018) using blastp. Sequences with an e-value < 1 × 10^-3, identity >25%, and coverage >75% were considered as potential homologs. On this basis, we found that 20 ChECs were “species-specific” with no homolog in other Colletotrichum species, 30 were “genus-specific” with homolog(s) limited to the genus Colletotrichum and 11 were “non-specific” with homolog(s) in other fungal genera (Supplementary Table 2). For the latter category, homologs were found most often in Fusarium oxysporum, Ceratocystis fimbriata and in species of Diaporthe and Rhynchosporium.

Expression profiles of the selected ChEC genes across four fungal developmental stages are presented in Figure 1B, based on mapping previous RNA-Seq data (O’Connell et al., 2012) to a revised assembly and annotation of the C. higginsianum genome (Dallery et al., 2017). Approximately two-thirds of the ChEC genes appear to be plant-induced, because they showed minimum expression in appressoria formed in vitro. Consistent with our previous qRT-PCR analysis of 17 ChECs (Kleemann et al., 2012), the 61 genes selected for study here also showed highly stage-specific expression. Thus, while 21 genes were preferentially expressed in appressoria (‘wave 1’), only eight were induced in both appressoria and the biotrophic phase (‘wave 2’), and the largest group (32 genes) were preferentially expressed during the biotrophic phase (‘wave 3’).

The predicted cDNA sequences of the selected ChECs were synthesized and cloned into a Gateway entry vector. In each case, the fungal signal peptide was omitted from the 5′ end, while adding an artificial start codon and retaining the original stop codon. The cloned ChEC sequences were shuttled into a binary destination vector suitable for transient over-expression of the proteins in N. benthamiana leaves via Agrobacterium infiltration. The vector provided a fusion to GFP at the N-terminus of each protein and expression was driven from the 35S promoter. After 2–3 days, the Agro-infiltrated tissues were examined by confocal microscopy to determine the subcellular localization of the fusion proteins.

Among the 61 GFP-tagged ChECs screened in this assay, 40 showed no specific localization in that they diffused throughout the plant nucleoplasm and cytoplasm, a pattern that was indistinguishable to that of GFP expressed alone (Figure 1C and Supplementary Figure 1). However, the remaining 21 fusion proteins were targeted either specifically or preferentially to particular compartments of the plant cell. Thus, nine GFP-ChECs were targeted to the plant nucleus, of which eight preferentially accumulated in the nucleolus, six labeled small punctate structures that were likely to be plant organelles, while three others decorated filamentous structures that resembled elements of the plant cytoskeleton (Figure 1C). A further three ChECs were excluded from the plant nucleus and confined to the plant cytoplasm (Figure 2). Their exclusion from the nucleus appears unrelated to their size because the predicted molecular weights (MW) of the mature proteins (ChEC21a = 10.38 kDa, ChEC30 = 16.78 kDa, ChEC103 = 6.99 kDa) were only slightly greater, or less than, the mean ChEC MW (11.45 kDa, Supplementary Table 1). In the case of ChEC30 and ChEC103, this localization was consistent with the detection of a putative nuclear export signal using the NetNES prediction tool (La Cour et al., 2004).

The subcellular localizations determined experimentally by GFP-tagging the ChECs were compared with localizations predicted by the LOCALIZER tool (Sperschneider et al., 2017) running in ‘effector mode’ or by WoLF PSORT (Horton et al., 2007) running in ‘plant mode’ after removing the fungal secretion signal. The results are compiled in Supplementary Table 1. LOCALIZER correctly predicted eight ChECs to be nuclear-targeted but eight others were false positives and one other was not predicted to be nuclear (false negative). LOCALIZER is currently not able to predict Golgi, peroxisomes or cytoskeleton as protein destinations. WoLF PSORT correctly predicted six nuclear-targeted ChECs with seven false positives and one false negative. WoLF PSORT predicted a chloroplast localization for no fewer than 15 of the ChECs (all incorrect based on our experimental results), while Golgi and cytoskeleton were not predicted as destinations for any of the proteins.

The integrity of the GFP fusion proteins was verified by Western blotting using anti-GFP antibodies. For all the ChECs showing a specific subcellular localization (21), we were able to confirm the presence of the full-length fusion protein without detectable free GFP, suggesting that cleavage of the FP from the effector had not occurred. We likewise verified the integrity of six GFP-ChECs that showed a nucleo-cytoplasmic localization, again confirming that their distribution was not due to cleavage and subsequent diffusion of free GFP (Supplementary Figure 2).

Nine ChECs were localized in the nuclei of N. benthamiana cells. To confirm this localization in the true host of C. higginsianum, we transiently expressed the proteins as N-terminal fusions with mRFP in transgenic A. thaliana seedlings stably expressing GFP fused to β-glucuronidase (GUS) and an NLS, which accumulates specifically in nuclei (Chytilova et al., 1999). All nine effector candidates that were imported into N. benthamiana nuclei also accumulated in A. thaliana nuclei (Figure 3). To further support these in situ localization data, we used four different computational algorithms to search for the presence of putative nuclear localization signals (NLS) within the predicted sequences (minus signal peptide) of these proteins, namely NLSpredict (Yachdav et al., 2014), cNLS Mapper (Kosugi et al., 2009), NLStradamus (Nguyen Ba et al., 2009), and WoLF PSORT (Horton et al., 2007). All nine of the nuclear-targeted ChECs possessed an NLS that was predicted by two or more different algorithms (Supplementary Table 3). In contrast, among the remaining 52 ChECs that did not accumulate in plant nuclei, only four were predicted to contain an NLS by two or more algorithms (ChEC9, ChEC51, ChEC96, and ChEC113).

Interestingly, the precise distribution of these nine ChECs in plant nuclei was not identical, and we were able to distinguish several different patterns. ChEC4 labeled the nucleoplasm but was excluded from the nucleolus in both N. benthamiana and A. thaliana (Figure 3). ChEC118 did not label the N. benthamiana nucleoplasm uniformly, instead showing a granular pattern reminiscent of chromatin structure (Figure 3). When GFP-ChEC118-labeled nuclei were in addition stained with DAPI, we observed that some regions of the nucleoplasm that were enriched with GFP-ChEC118 were not stained by DAPI (Supplementary Figure 3). This partitioning of the plant DNA contrasts to normal N. benthamiana nuclei at interphase, where DAPI labels the nucleoplasm uniformly (Supplementary Figure 4). However, this partitioning of the nucleoplasm was not visible in nuclei of A. thaliana cells expressing RFP-ChEC118 (Figure 3).

Six other nuclear-targeted proteins, namely ChEC74, ChEC98, ChEC104, ChEC106, ChEC108, and ChEC111, were concentrated inside the nucleolus and other sub-nuclear compartments in both N. benthamiana and A. thaliana (Figure 3). Punctate accumulations of ChEC98 of varying size were also present on the surface of nuclei in both plant species. These structures do not resemble known compartments of the plant cell and may represent artefactual protein aggregates caused by over-expression of the fusion protein. ChEC74 and ChEC104 had similar localization patterns in that they preferentially labeled nucleoli more than the nucleoplasm, and both were predicted to harbor a nucleolar targeting signal using the NOD predictor (Scott et al., 2010, 2011). In addition to being concentrated in the nucleolus, ChEC108 also labeled smaller sub-nuclear structures that resembled Cajal bodies in size (0.2–2 μm) and number (1–6 per nucleus) in both N. benthamiana and A. thaliana (Figure 3). In order to identify these compartments, we co-expressed GFP-ChEC108 or RFP-ChEC108 together with protein markers labeling the nucleolus and Cajal bodies, namely Fibrillarin 2 (RFP-FIB2) and U2 Small nuclear ribonucleoprotein B (GFP-U2B; Shaw et al., 2014). A sub-set of small ChEC108-positive structures co-localized with FIB2 or U2B, and are therefore likely to be Cajal bodies (Supplementary Figure 5). However, other sub-nuclear compartments were only labeled by ChEC108 and therefore remain unidentified.

Remarkably, and in contrast to all the other nuclear-targeted ChECs, transient over-expression of GFP-ChEC106 in N. benthamiana resulted in an increase in the size of the nucleus in transformed cells, involving an inflation of the nucleoplasm but not the nucleolus (Figure 3 and Supplementary Figure 4). Thus, the size of nuclei in cells expressing GFP-ChEC106 (mean area = 175.1 μm², n = 106) was 2.2-fold larger than in cells expressing GFP-ChEC104 (mean area = 80.0 μm², n = 49) and 2.9-fold larger than in untransformed cells, in which nuclei were stained with DAPI (mean area = 59.6 μm², n = 81) (Supplementary Figure 4D). The observed differences were highly significant in all pair-wise comparisons (Mann–Whitney non-parametric test, p < 0.01). The nuclei of transformed cells did not appear enlarged in A. thaliana seedlings expressing RFP-ChEC106 (Figure 3), although nuclear sizes were not quantified.

Six of the GFP-ChECs became concentrated in small punctate structures moving rapidly in the plant cytoplasm, which could represent plant organelles. None of these were chloroplasts, which could be easily distinguished by their larger size and chlorophyll autofluorescence. To identify the labeled compartments, we performed co-localization experiments by transiently co-expressing the GFP-ChECs in N. benthamiana cells together with organelle-specific markers tagged with red FPs. Peroxisomes were labeled with PTS2-DsRed (peroxisome targeting signal 2 fused to DsRed; Fujimoto et al., 2009). Mitochondria were labeled with Mt-RFP (mitochondrial targeting signal of the Arabidopsis ATPase δ-subunit fused to mRFP (Arimura and Tsutsumi, 2002). The trans-Golgi cisternal membranes of Golgi stacks were labeled with ST-mRFP (signal anchor of rat sialyltransferase; Sparkes et al., 2006; Uemura et al., 2012). Early and late endosomes, respectively, were labeled using the Rab-like GTPases ARA6 and ARA7 fused to RFP (Ueda et al., 2001). Finally, the trans-Golgi network/early endosome compartment was labeled with the syntaxin Syp61 fused to RFP (Choi et al., 2013). Among the GFP-ChECs labeling putative plant organelles, ChEC36 and ChEC39 did not co-localize with any of the six organelle markers and labeled punctate structures of two different sizes (mean diameter 0.98 μm and 2.15 μm for ChEC36 and ChEC39, respectively, n = 36) (Supplementary Figure 6).

GFP-ChEC21 perfectly co-localized together with ST-mRFP, consistent with this protein being associated with plant Golgi stacks (Figure 4). At higher magnification, the organelles labeled by GFP-ChEC21 and ST-mRFP could be resolved as distinct ring-shaped structures with a bright periphery and a dark core (Figure 4), as reported previously for Golgi stacks labeled by ST-GFP and cellulose synthase CESA3-GFP (Boevink et al., 1998; Crowell et al., 2009). This characteristic doughnut-like labeling pattern suggests that ChEC21 labels the rim of plant Golgi stacks.

Three other GFP-tagged proteins, namely ChEC51a, ChEC89, and ChEC96, labeled small highly mobile organelles of similar size (Figure 5A). All three fusion proteins co-localized with PTS2-DsRed, suggesting that the punctate structures labeled by these ChECs are plant peroxisomes (Figure 5B). While ChEC89 filled the entire peroxisome matrix, ChEC51a and ChEC96 were concentrated into a smaller punctate structure that appeared to be located inside the peroxisome (Figure 5B). To examine the localization of GFP-ChEC89 and GFP-ChEC96 at higher resolution, we used transmission electron microscopy and immunogold labeling with antibodies specific for RFP to detect the PTS2-DsRed peroxisome marker, and antibodies specific to GFP to label the ChECs. In adjacent ultrathin sections through the same peroxisome, both GFP-ChEC89 and PTS2-DsRed were detected in the peroxisome matrix, with little or no labeling detectable on a large electron-opaque inclusion within the matrix, which probably corresponds to the catalase crystal (Figures 6A,B). In contrast, GFP-ChEC96 appeared less uniformly distributed through the peroxisome matrix than either GFP-ChEC89 or PTS2-DsRed, and in some sections the GFP-ChEC96 labeling could be seen concentrated into a small area of the peroxisome matrix (Figures 6C,D). This is unlikely to represent the catalase crystal because in those peroxisomes where the crystal was clearly recognizable, it showed little or no labeling with anti-GFP antibodies (Figures 6E,F). These immunolabeling experiments therefore confirmed that both ChEC89 and ChEC96 are imported into the peroxisome matrix and that ChEC96, but not ChEC89, becomes concentrated into small inclusions that are distinct from the catalase crystal. Similar immunolocalization experiments were not preformed with ChEC51a because the transient over-expression of this protein in N. benthamiana was found to cause necrosis in a subset of the transformed cells.

The import of proteins from the cytosol into the peroxisome matrix is mediated by conserved peroxisome targeting signals, either of type 1 (PTS1) or type 2 (PTS2), which can be predicted using computational tools (Lingner et al., 2011). The plant-specific PredPlantPTS1 algorithm (Reumann et al., 2012) predicted the presence of a PTS1 tripeptide at the C-terminus of ChEC51a (SKL, 100% targeting probability) but not in ChEC96 (PRL, 8.7% targeting probability) (Figure 7A). Nevertheless, SKL and PRL are both canonical PTS1 signals found in most eukaryotes that have been shown to function in plants (Lingner et al., 2011; Reumann et al., 2012). PredPlantPTS1 takes into account not only the C-terminal tripeptide but also the preceding 11 amino acids, and the negative prediction for ChEC96 reflects the presence of residues that potentially inhibit peroxisome targeting (Reumann et al., 2012). However, ChEC96 was predicted to contain a PTS1 signal (56.7% targeting probability) by a fungi-specific prediction model (Nötzel et al., 2016). The protein sequence of ChEC89, which is less than half the length of ChEC51a and ChEC96 (Figure 7A), was not predicted to contain either PTS1 or PTS2 signals using PredPlantPTS1, PTS1Prowler (Bodén and Hawkins, 2005) or PeroxisomeDB (Schlüter et al., 2010).

To experimentally verify the functionality of the C-terminal tripeptides in ChEC51a and ChEC96, we deleted the corresponding nucleotide sequences and added an artificial stop codon. The transient expression of these truncated proteins showed that they were no longer targeted to plant peroxisomes and instead showed a uniform nucleo-cytoplasmic distribution (Figure 7B). Based on these experiments, we conclude that both ChEC51a and ChEC96 harbor functional PTS1 tripeptides that are required to permit their translocation into plant peroxisomes via the classical PTS1 pathway.

Three effector candidates, namely ChEC9, ChEC17, and ChEC113, labeled arrays of long filamentous structures in the cortical cytoplasm of N. benthamiana cells which resembled elements of the plant cytoskeleton (Figure 8). GFP-ChEC9 exclusively labeled these cytoskeleton filaments (Figure 8A) whereas GFP-ChEC17 was also concentrated in granular, punctate structures located along the filaments and inside plant nuclei (Figures 8B,C). However, no NLS was detected in the protein sequence of ChEC17 using NLSpredict, cNLS Mapper, NLStradamus, or WoLF PSORT. Similar to GFP-ChEC9, GFP-ChEC113 also labeled small punctate structures that were closely associated with the cytoskeleton filaments, but nuclei were not labeled by this fusion protein (Figure 8D).

In order to distinguish whether these proteins were associated with plant microtubules or actin microfilaments, we either transiently expressed them as N-terminal RFP fusions in transgenic N. benthamiana plants (line CB13) stably expressing tubulin alpha 6 (TUA6) fused to GFP (Ueda et al., 1999; Gillespie et al., 2002), or transiently co-expressed them as N-terminal RFP fusions together with the fimbrin actin-binding domain 2 (FABD2) fused to GFP (Sheahan et al., 2004; Voigt et al., 2005). In these experiments, we found that ChEC9, ChEC17, and ChEC113 were all perfectly co-localized with GFP-TUA6-labeled microtubules in every N. benthamiana cell examined (Figure 9A). In contrast, the RFP-tagged ChECs displayed no detectable co-localization with actin microfilaments (Figure 9B). Consistent with this microtubule localization, ChEC113 was predicted to be a microtubule-associated protein using the MAPanalyzer tool (Zhou et al., 2015); however, ChEC9 and ChEC17 were not.