To better understand the timeframe during which the auxin receptor family diversified, we built upon previous phylogenetic analyses (Parry et al., 2009; Mutte et al., 2018) with more taxon sampling at key nodes. As shown earlier (Hori et al., 2014; Mutte et al., 2018), the TIR1/AFB genes likely evolved from a gene encoding an F-Box/LRR protein similar to those present in the genomes of extant streptophyte algae. These algal proteins form a sister clade to three distinct land plant F-Box families, the TIR1/AFB auxin receptors, the COI1 jasmonate-Ile (or dinor-OPDA) receptors, and the ‘XFB’ clade of unknown function conserved in the genomes of mosses and some lycophytes but not in other land plants (Prigge et al., 2010; Bowman et al., 2019). While the last common ancestors of land plants and of vascular plants had only one TIR1/AFB gene, three clades of auxin receptors were established prior to the radiation of euphyllophytes (ferns plus seed plants) over 400 million years ago (Morris et al., 2018; Figure 1—figure supplement 1A). Another gene duplication event prior to angiosperm radiation split the TIR1/AFB1 clade from the AFB2/AFB3 clade. Receptors from each of the four clades are not retained in the genome of every flowering plant. For example, AFB6 orthologs are not present in the genomes of core Brassicales species—including Arabidopsis—nor those of Poaceae species including rice and maize. The gene duplication event establishing the distinct TIR1 and AFB1 clades is coincident with the At-β WGD event at the base of Brassicales, while both the AFB2/AFB3 and the AFB4/AFB5 duplication events coincide with the more recent At-α WGD prior to divergence of the Brassicaceae family (Figure 1—figure supplement 1A; Schranz and Mitchell-Olds, 2006).

One noteworthy aspect of the phylogenetic tree is the pronounced branch-length asymmetry within the TIR1+AFB1 clade (Figure 1—figure supplement 1). Since the last common ancestor of Arabidopsis (Brassicaceae) and Tarenaya (Cleomaceae), the AFB1 gene has accumulated over three times as many non-synonymous changes as TIR1 (Figure 1—figure supplement 1B) despite being under selection based on the ratio of non-synonymous and synonymous substitutions (Delker et al., 2010; Wright et al., 2017). AFB1 also differs from the other TIR1/AFBs in that it contains two of three substitutions in the first α-helix of the F-Box that were each previously shown to weaken TIR1’s interaction with CUL1 (Yu et al., 2015). The substitution with the largest effect, Glu8Lys (equivalent to Glu12Lys in TIR1), appeared shortly after the At-β WGD that produced AFB1, and the Phe14Leu substitution appeared prior to the crown group of the Brassicaceae family (Figure 1—figure supplement 1C). Interestingly, AFB1 orthologs from members of the Camelina genus—C. sativa (all three homeologs), C. laxa, C. hispida, and C. rumelica—additionally contain the third substitution (Figure 1—figure supplement 1C).

Previous studies have assessed the functional overlap between the TIR1, AFB1, AFB2 and AFB3 genes (Dharmasiri et al., 2005; Parry et al., 2009) and separately between the AFB4 and AFB5 genes (Prigge et al., 2016). To study the genetic interactions between all members of the family, and to determine the effects of the complete absence of TIR1/AFB-mediated auxin signaling, lines with strong loss-of-function mutations in the six TIR1/AFB genes were intercrossed to generate all sixty-three mutant combinations. We used the following alleles tir1-1, afb1-3, afb2-3, afb3-4, afb4-8, and afb5-5 (Figure 1—figure supplement 2A; (Ruegger et al., 1998; Parry et al., 2009; Prigge et al., 2016). The tir1-1 allele, which causes an amino acid substitution within the leucine-rich repeat domain of the protein, has been reported to act as a dominant-negative allele (Dezfulian et al., 2016; Wright et al., 2017). However, we found that the root elongation phenotype of plants heterozygous for the tir1-1, tir1-10, and tir1-9 alleles were not significantly different from each other and each displays a semi-dominant phenotype (Figure 1—figure supplement 2B). These results argue against a dominant negative effect for tir1-1 since neither tir1-9 or tir1-10 produce detectable levels of transcript (Ruegger et al., 1998; Parry et al., 2009). Nevertheless, because it is possible that a dominant-negative effect might be revealed in higher-order mutants and because the afb2-3 allele may not be a complete null allele (Parry et al., 2009), we generated selected mutant combinations also using the tir1-10 (Parry et al., 2009) and the afb2-1 (Dharmasiri et al., 2005) T-DNA insertion alleles. The afb2-1 allele was introgressed from the Ws-2 background into the Col-0 background through at least eight crosses. For brevity, mutant line names will be simplified such that ‘tir1afb25’ corresponds to the tir1-1 afb2-3 afb5-5 triple mutant line, for example, unless other allele numbers are specified.

The sixty-three mutant combinations displayed a wide range of phenotypes from indistinguishable from wild type to early-embryo lethality (Figure 1—figure supplement 3; Supplementary file 1). The non-lethal higher-order mutant combinations displayed a cohort of phenotypes associated with mutants defective in auxin signaling: smaller rosettes, reduced inflorescence height, reduced apical dominance, fewer lateral roots, and partially or wholly valveless gynoecia (Figure 1; Figure 1—figure supplement 3; Supplementary file 1). The three viable quintuple mutants—tir1afb1245, tir1afb1345, and afb12345—had rosettes approximately half the diameter and inflorescences less than half the height of WT Col-0. Despite being smaller, these lines produced approximately twice as many branches as WT (Figure 1A; Figure 1—figure supplement 3; Supplementary file 1). Remarkably, lines retaining only one copy of TIR1 (tir1/+ afb12345) or one copy of AFB2 (afb2/+ tir1afb1345) were viable. The rosettes of these two lines were much smaller than those of WT plants with the tir1/+ afb12345’s rosette phenotype being slightly more severe (Figure 1B). In contrast, afb2/+ tir1afb1345 plants developed shorter primary inflorescences and appeared to completely lack apical dominance as all axillary meristems became active upon flowering. The afb2/+ tir1afb1345 and tir1/+ afb12345 plants rarely produced seeds. Lines containing the alternate alleles—afb2-1/+ tir1-10 afb1345 and tir1-10/+ afb2-1 afb1345—displayed phenotypes indistinguishable from the corresponding lines (Figure 1B).

Auxin plays an important role in many aspects of root development. To begin to assess the role of the TIR1/AFBs during root growth, we measured the effect of exogenous IAA on primary root growth in the mutant lines. The responses ranged from indistinguishable from WT to nearly insensitive to 0.5 µM IAA (Figure 1—figure supplement 3E), where the roots of lines containing the tir1 and afb2 mutations displayed strong IAA resistance (Dharmasiri et al., 2005; Parry et al., 2009). In addition, we found that the afb3 and afb5 mutations had substantial effects on auxin response, while the afb4 mutation had a more modest effect. The mutant lines also responded similarly to exogenous auxin with respect to lateral root production. The lines more resistant to IAA in the root elongation assay tended to produce fewer lateral roots (Figure 1—figure supplement 3D and E).

Each of the tir1/afb mutations, except for afb1, affected the above-described phenotypes but to varying extents. To appraise the effects of each mutation on several plant phenotypes, we plotted the mean values for each phenotype minus that of the corresponding line without that mutation. Larger effects are indicated by greater deviations from zero. For both the root elongation assay and the induction of lateral root primordia, the tir1 allele had the largest effect with the afb2, afb5, afb3, and afb4 mutations having smaller median effects (Figure 2A and B). The afb1 mutation had little or no effect on root elongation but, surprisingly, had an opposite effect on lateral root formation.

Only tir1 and, to a lesser degree, afb2 affect rosette diameter in most contexts with the median effects for afb3, afb4, and afb5 being very close to zero (Figure 2C). However, they have huge collective effects, where the afb2345 quadruple mutant is over 6 cm smaller than each of the four triple mutants (blue arrowheads; Figure 2C). Consistent with previous reports that AFB5 plays a key role in regulating inflorescence branching and height (Prigge et al., 2016; Ligerot et al., 2017), the afb5 mutation has the largest effect on these phenotypes, although each mutation, except for afb1, had some effect (Figure 2D and E).

While the afb1 mutation had minimal effect on most aspects of plant growth, it suppressed the lateral root phenotype of some mutant lines both with and without auxin treatment (Figure 2B; below). We found that the afb1 mutation suppressed the phenotype of both the afb234 (2.15 ± 0.13 versus 1.75 ± 0.10 lateral roots/cm) and afb345 triple mutants (3.10 ± 0.13 versus 1.96 ± 0.14 lateral roots/cm) measured after 12 days on media not supplemented with IAA. This behavior was not observed in an otherwise WT background (2.76 ± 0.11 for afb1 versus 3.23 ± 0.15 lateral roots/cm for Col-0) nor in a tir1-1 background (1.75 ± 0.08 versus 2.34 ± 0.09 lateral roots/cm for tir1). Each of the pairs were significantly different (two-tailed t-test, p<0.03).

The tir1afb23 and tir1afb123 lines were previously shown to display a variably penetrant phenotype in which seedlings lack roots, lack both roots and hypocotyls, or fail to germinate (Dharmasiri et al., 2005; Parry et al., 2009). All lines homozygous for both tir1 and afb2 plus either afb3, afb4, or afb5 display these defects to some degree ranging from 1% in tir1afb24% to 99% in tir1afb1234 (Supplementary file 1; Figure 1—figure supplement 4).

We had noticed a sizeable difference in the proportion of seedlings lacking roots from different batches of seeds. To test whether the temperature at which the seeds mature affects the penetrance of the rootless seedling phenotype, we grew tir1afb23, tir1afb123, tir1afb245 and WT in parallel at 17°C, 20°C, and 23°C and scored the progeny seedling phenotypes (Figure 1—figure supplement 4). The penetrance of the phenotype for all three lines was significantly lower at 20°C than at either 17°C or 23°C for all with the exception that the difference with tir1afb245 at 17°C was not significant using the Fisher’s exact test. This suggests that aspects of the auxin regulatory system are sensitive to temperature.

Because tir1afb235 seedlings were not identified among the progeny of tir1/+ afb235 or afb2/+ tir1afb35 plants, we examined developing embryos dissected from the siliques from these lines. Eleven of 41 (27%) and 52 of 213 (24%), respectively, of the embryos from each line lacked cotyledons and had over-proliferated suspensors while the rest had a WT phenotype (Figure 3A, A').

Because all mutant combinations expected to produce one-quarter tir1afb2345 progeny were either seedling lethal or infertile, we created a transgene that hemizygously complements these phenotype and segregates as a single locus. We assembled the complementing genomic fragments encoding TIR1, AFB2, and AFB5, each carboxy-terminally fused with the coding sequences for different monomeric fluorescent proteins (mOrange2, mCitrine, and mCherry, respectively) concatenated into a single binary plasmid (Figure 3—figure supplement 1). This construct was transformed into progeny of tir1/+ afb5/+ afb1234 plants, backcrossed and selfed to obtain a sextuple mutant background, and two TIR1/AFB5/AFB2 lines were identified that complemented the sextuple mutant phenotype when hemizygous and segregated as a single locus. Using this approach, one-quarter of the progeny of plants hemizygous for these transgenes display embryo defects, while the complemented siblings are easily identified because they expressed fluorescent TIR1, AFB2, and AFB5 fusion proteins.

The earliest potential difference between sextuple mutants and the complemented siblings is that the initial division of the embryo proper was occasionally displaced from vertical in sextuple 2-cell embryos (3 of 19 were >12° from vertical) compared to complemented sibling embryos (0 of 64), however the average angles from vertical were not significantly different (p=0.32) (Figure 3B–C versus 3B′–C′, 3N). The third round of divisions in the embryo proper separates the upper and lower tiers with the lower surface of the upper tier typically being slightly convex, and this curvature is significantly more prominent in the sextuple mutants (p=1.4 × 10⁻⁷; Figure 3D versus 3D′, 3O). Later, during the transition from the 8-cell to the 16-cell embryos, nearly all cell divisions in the complemented embryos are oriented periclinally, as in WT embryos. In contrast, 69% of these division are anticlinal in the mutant embryos (Figure 3E and E', 3P). In WT 32-cell stage embryos, the hypophysis cell normally divides asymmetrically to produce the lens-shaped cell which is required for the formation of the embryonic root. This division was delayed in the mutant, and when it occurred, was symmetrical (Figure 3G–H and 3G′–H′). Later, the cells of the embryo proper slow or cease dividing and the cells of the suspensor begin to proliferate and invariably produce a radially symmetric terminal phenotype (Figure 3I–M, 3I′–M′). Around the stage where complemented siblings are at the bent-cotyledon stage, the cells of the sextuple mutant senesce and seed development is aborted (Figure 3—figure supplement 2). Hence, the sextuple mutant reveals the importance of TIR1/AFB auxin response machinery from the earliest stages of embryogenesis.

To learn more about the early embryo defects, we introgressed marker genes into lines segregating the tir1afb235 quadruple mutant. This quadruple mutant displays a phenotype indistinguishable from that of the sextuple mutant from at least the dermatogen stage, when quadruple mutants can first be reliably distinguished from non-quadruple mutants (Figure 4). Expression of the auxin-responsive marker DR5_rev:3×Venus-N7 was undetectable in embryos displaying the mutant phenotype (Figure 4E–G; compare to Figure 4A–C; 0/10 in quadruple mutants and 26/26 in non-mutant siblings) indicating that auxin-regulated transcription through the remaining receptors, AFB4 and AFB1, is minimal at most during embryogenesis. Similarly, the quiescent center marker WOX5:GFP was not detected in the mutant embryos (Figure 4H), whereas in wild type the reporter is first expressed in the hypophysis prior to its asymmetric division then persists in the quiescent center cells (Figure 4D; 0/11 in quadruple mutants and 28/28 in non-mutant siblings).

Auxin efflux reporter PIN7-Venus is normally expressed in the suspensor, hypophysis, hypophysis-derived cells, and weakly in protodermal cells of the lower tier (Figure 4K–L). In mutant embryos, PIN7-Venus is faintly detectable in these cells in globular-stage embryos. Unexpectedly, the signal is much stronger in protodermal cells of the embryo proper, especially in the lower tier, by the 32-cell stage (Figure 4O–P; 7/8 quadruple mutants and 0/14 siblings). The same pattern was observed with the PIN7-GFP marker (Figure 4U; 8/8 quadruple mutants and 0/30 siblings). The auxin efflux reporter PIN1-Venus is initially expressed in a reciprocal pattern to PIN7-Venus, in all the cells above the hypophysis except the lower-tier protodermal cells and is later refined to strips from the provascular cells out to the cotyledon tips (Figure 4I–J; 12/12 of phenotypically normal embryos). In the mutants, PIN1-Venus signal is reduced and restricted primarily to apical protodermal cells (Figure 4M–N; 9/9 quadruple mutants). The NTT-YPet marker gene is normally first strongly expressed in 8- to 16-cell embryos in the nuclei of suspensor cells and the hypophysis and persists in the suspensor and the hypophysis-derived cells in later embryo stages (Figure 4R–T; 58/59 phenotypically normal embryos) (Crawford et al., 2015). In mutants, NTT-YPet appears normally in most suspensor cells, but not always including the hypophysis (2/6 32-cell quadruple mutant embryos had signal above background in the hypophysis), and is progressively lost in the distal suspensor cells before the abnormal lateral cell divisions occur (Figure 4V–X; 4/4 late-globular-stage mutants lacked signal in both the hypophysis and the adjacent suspensor cell). This is very similar to NTT-YPet expression in a monopteros mutant embryos (Crawford et al., 2015).

Because the maternal supply of auxin and the endosperm both play important roles in embryo development, it is possible that female gametophytes lacking auxin receptors would not be viable. Although the incidence of sextuple mutant embryos shows that gametophytically-expressed auxin receptors are not required for viability, it is possible that they contribute to robust transmission. To test the transmission through sextuple mutant megagametophytes and pollen, we carried out reciprocal crosses between wild type (Col-0) and the hemizygously TIR1/AFB5/AFB2-complemented sextuple mutant. If the sextuple mutant gametophyte survives and is fertilized, the progeny’s embryo lethality would be rescued by wild-type copies of each receptor provided by the Col-0 parent. The F₁ progeny were scored for the presence of the transgene to infer the sextuple’s transmission rates through both gametophytes (Supplementary file 2). The sextuple mutant was transmitted nearly as well as without the complementing transgene as with it through both the pollen (49.4%) and the female gametophyte (47.9%) (χ² test p=0.81 and 0.43, respectively). This indicates that gametophytically expressed TIR1/AFBs do not contribute to gametophytic viability.

To reveal whether differences in expression pattern can account for the relative importance of the TIR1/AFBs in different aspects of growth and development, C-terminal fusions with the bright, relatively fast-maturing, monomeric fluorescent protein mCitrine were produced for each TIR1/AFB protein in the corresponding single mutant background. Each transgene complemented the mutant phenotypes (Figure 5—figure supplement 1). The fluorescent signal in the AFB5-mCitrine lines was fairly uniformly distributed in shoot apices (Figure 5F), while in the AFB3-mCitrine, AFB2-mCitrine, and TIR1-mCitrine lines, fluorescence was more restricted to young primordia and meristem peripheral zones (Figure 5A and C–D). Within organ primordia, TIR1-mCitrine appears to be strongest in the adaxial domains of the youngest primordia. Signal for the AFB4-mCitrine line was barely detectable (Figure 5E), while that of AFB1-mCitrine was very strong and largely complementary to TIR1-mCitrine in that the strongest signal was in abaxial domains and in the stem (Figure 5B). The expression patterns in either primary or lateral roots for each TIR1/AFB gene except AFB4 translationally fused to a YFP have been reported previously (Prigge et al., 2016; Wang et al., 2016; Rast-Somssich et al., 2017; Roychoudhry et al., 2017). TIR1-, AFB2-, AFB3-, and AFB5-mCitrine signal was uniformly detected throughout the root meristematic region and fainter signal detected in the root cap cells (Figure 5G, I–J and L; Figure 5—figure supplement 2) as shown previously. AFB1-mCitrine is very highly expressed throughout the root except for the columella, cortex, endodermis, and pericycle of the meristematic region (Figure 5H). The expression pattern of an AFB4-mCitrine line (line #3), hypersensitive to the synthetic auxin picloram, was comparable to that of AFB5-mCitrine while that of AFB4-mCitrine line #1 that complemented the afb4 phenotype was barely detectable (Figure 5K; Figure 5—figure supplement 1D, Figure 5—figure supplement 2I–L). In embryos, TIR1-, AFB2-, AFB3-, and AFB5-mCitrine accumulate fairly uniformly throughout the embryos and suspensors while AFB4-mCitrine’s signal was close to background levels and AFB1-mCitrine was undetectable (Figure 5T–Y).

The subcellular localization of different TIR1/AFB proteins varied substantially (Figure 5M–S; Figure 5—figure supplement 3). We quantified this variation more precisely by measuring the relative level of each protein in the nucleus versus outside the nucleus in epidermal cells of the root elongation zone based on mCitrine fluorescence (Figure 5S). TIR1-mCitrine is primarily in nuclei while significant amounts of AFB2 through AFB5 are present in the cytoplasm. Strikingly, AFB1-mCitrine appears primarily outside the nuclei. The localizations are consistent across multiple lines and, when tested, different fluorescent protein tags (Figure 5—figure supplement 2). Hence, despite being from the same clade, TIR1 and AFB1 proteins exhibit contrasting patterns of primarily nuclear and cytoplasmic subcellular localizations, respectively.

Gravitropic curvature of the root is a rapid auxin-regulated growth response that requires asymmetric distribution of auxin between the upper and lower side of the root (Sato et al., 2015). According to the current model, auxin has two modes of action during gravitropism: a rapid nongenomic phase, followed by a transcriptional phase that is dependent on the TIR1/AFB proteins (Shih et al., 2015). Surprisingly, recent studies demonstrate that rapid, nongenomic auxin inhibition of root growth is dependent on the TIR1/AFBs (Fendrych et al., 2018). To determine the relative contribution of the TIR1/AFB family members to the rapid response, we measured the effect of 10 nM IAA on root growth in various tir1/afb lines over a 20 min time period (Figure 6; Figure 6—figure supplement 1; Figure 6—videos 1–2). The results in Figure 6 show that each of the TIR1/AFBs contributes to rapid root growth inhibition but that, surprisingly, the afb1-3 mutant is almost completely resistant to auxin indicating that AFB1 is the dominant auxin receptor for this response. Expression of AFB1-mCitrine under control of the AFB1 promoter restored the wild-level of auxin response. The behavior of the afb1 mutant is particularly remarkable since the mutant is not affected in any of the other auxin-regulated growth processes that we characterized, with the possible exception of lateral root formation. This includes long-term inhibition of root growth. Thus, the afb1 mutant is a useful tool to discriminate between nongenomic and transcriptional auxin responses.

As a contrast we also examined the effect of auxin on etiolated hypocotyl growth in the mutant lines. This response is slower than the root response and depends on the canonical nuclear TIR1/AFB pathway (Fendrych et al., 2016). Dissected hypocotyl segments from etiolated seedlings were treated with 5 µM NAA and imaged every 10 min for 180 min. The response of the mutant lines was complex (Figure 6—figure supplement 2). Several lines were clearly resistant to auxin, particularly tir1afb1245 and tir1afb245. Notably, comparison of these lines suggests that the afb1 mutation did not contribute to resistance. Other lines also lacking both AFB4 and AFB5—afb1345, tir1345, afb1245, and afb12345—display a moderate level of resistance. Finally, three lines, tir1afb134, tir1afb23, and tir1afb124 are hypersensitive to auxin in this assay (Figure 6—figure supplement 2).

Gravitropic root curvature is first apparent less than 10 min after a gravity stimulus (Shih et al., 2015). It has been proposed that the early stage of gravitropism is mediated by a non-genomic auxin response while prolonged root curvature requires auxin regulated transcription (Sato et al., 2015). Since AFB1, and to a lesser extent, the other TIR1/AFBs, contribute to nongenomic inhibition of root elongation, we wondered if they are required for the early gravitropic response. To test this possibility, we performed gravitropism assays on a number of tir1/afb lines (Figure 7; Figure 7—figure supplement 1). The gravitropic response can be divided into three phases. A slow or lag phase which occurs over the first 90 min in Col-0, followed by a 3 hr linear phase and finishing with a plateau phase. The tir1afb345 line exhibits a slower gravitropic response during the linear phase and a reduced angle at plateau compared to WT (Figure 7A) as expected based on results with other tir1/afb mutant combinations (Dharmasiri et al., 2005). Strikingly, the tir1afb1345 mutant exhibited an additional decrease in the gravitropic response during the initial lag phase demonstrating that AFB1 is required for this phase (Figure 7A; Figure 7—figure supplement 1A). The tir1afb1 line showed a similar decrease in the lag phase compared to tir1 (Figure 7B; Figure 7—figure supplement 1B). It seems likely that TIR1 also contributes to the early response since the tir1afb1 double mutant showed a stronger delay compared to either single mutant. In addition, both tir1 and afb1 displayed a reduced early response in one of the two experiments (Figure 7—figure supplement 1B). Since other members of the family also confer low levels of auxin resistance in the rapid root growth response, these proteins may also make a small contribution to the early phase of gravitropism (Figure 6A). Further, both afb1 AFB1-mCitrine lines responded appreciably faster than the afb1 mutant during the first 2 hr with the brighter of the two mCitrine lines, line #7, exhibiting a difference by 30 min (Figure 7C; Figure 7—figure supplement 1C). Interestingly, both mCitrine lines started to plateau earlier and at a reduced angle compared to wild type while afb1 plateaued later and at an increased angle, suggesting that AFB1 and the rapid response also play a role at later stages of the gravitropic bending response.

The sources for the amino-acid sequences (Figure 1—figure supplement 1A) and CDS (Figure 1—figure supplement 1C) are listed in Supplementary file 3 (Jiao et al., 2011; Goodstein et al., 2012; Johnson et al., 2012; Matasci et al., 2014; Wickett et al., 2014; Xie et al., 2014; One Thousand Plant Transcriptomes Initiative, 2019). Taxa were selected based on availability, quality, and diverse sampling at key nodes. A reduced set was included for COI1 homologs. The AFB1 genes from Camelina hispida, C. laxa, and C. rumelica were amplified from genomic DNA using Phusion Polymerase (New England Biolabs or ThermoFisher) and primers to regions of the 5′ and 3′ UTRs conserved in all three C. sativa AFB1 genes in the C. sativa genome (Kagale et al., 2014). The PCR products were subcloned, and three C. hispida and C. laxa clones and a single C. rumelica clone were sequenced. The CamhiAFB1 and CamlaAFB1 sequences included in analysis appeared in two of the three clones (GenBank accession numbers MK423960–MK423962).

To build the alignment of F-Box-LRR protein sequences, sequences from distinct subclades were aligned using T-COFFEE v11.00 (Notredame et al., 2000) to identify and trim unique unalignable regions from individual sequences before aligning the whole set. Ambiguous regions of the full alignments were removed in Mesquite v3.5 (Maddison and Maddison, 2018). The raw alignment of nucleotide CDS sequences of Brassicales TIR1/AFB1 genes was adjusted so that gaps fell between adjacent codons. Phylogenetic trees were inferred using MrBayes v3.2.6 (Ronquist et al., 2012). For the TIR1/AFB/XFB/COI1 phylogeny, a total of six runs of four chains were split between two Apple iMac computers using the parameters aamodelpr = mixed, nst = 6, and rates = invgamma. Only four of the six runs had converged after 16 million generations, so the analysis was restarted with three runs each starting with the best tree from one of the initial runs and with more heating (temp = 0.5) for 10 million generations. The TIR1/AFB1 nucleotide alignments were partitioned by codon position with ratepr = variable, nst = 6, rates = invgamma with three runs of 4 chains run for five million generations. The consensus trees were viewed using FigTree v1.4.4 (Rambaut, 2018).

The alleles used—tir1-1, tir1-9, tir1-10, afb1-3, afb2-1, afb2-3, afb3-1, afb3-4, afb4-8, and afb5-5—have been described previously (Ruegger et al., 1998; Dharmasiri et al., 2005; Parry et al., 2009; Prigge et al., 2016). Seeds from Camelina species were provided by the United States National Plant Germplasm System (USDA-ARS, USA): C. hispida (PI 650133), C. laxa (PI 633185), and C. rumelica (PI 650138). Unless noted, plants were grown at 22°C long-day (16:8) conditions on ½×Murashige and Skoog media with 0.8% agar, 1% sucrose, and 2.5 mM MES, pH 5.7, or in a 2:1 mixture of soil mix (Sunshine LC1 or ProMix BX) and vermiculite. Leaf DNA was isolated with a protocol adapted from Edwards et al. (1991) to use steel BBs (Daisy Outdoor Products), 2 ml microcentrifuge tubes, and 20-tube holders (H37080-0020, Bel-Art). See Supplementary file 4 for primers used for genotyping.

NTT-2×YPet (Crawford et al., 2015), PIN7-GFP (Blilou et al., 2005), DR5_rev:3×Venus-N7 (Heisler et al., 2005), WOX5:GFP_ER (Blilou et al., 2005). The recombineered PIN1-Venus and PIN7-Venus markers (Zhou et al., 2011) were obtained from the Arabidopsis Biological Resource Center (CS67184 and CS67186). Previously characterized PIN1-GFP lines could not be used because of tight linkage to AFB2 (CS9362) and co-segregation with Ler-derived enhancers of the afb2/+ tir1afb35 phenotype (CS23889). Each marker was introgressed into lines segregating the tir1afb235 quadruple mutant by two sequential crosses, PCR genotyping, and selfing. Marker line homozygosity was confirmed in F₁ seedlings from test crosses to WT. The UBQ10:H2B-mTurquoise2 marker was assembled by combining the pK7m34GW destination vector (Karimi et al., 2007), UBQ10prom_P4P1R (Jaillais et al., 2011) (provided by Nottingham Arabidopsis Stock Centre, N2106315), H2B_noStop/pDONR207 (provided by Frederic Berger), and 2×mTurqoise2/pDONR-P2RP3 using the LR Recombinase System (Life Technologies). For 2×mTurqoise2/pDONR-P2RP3, the mTurquoise2 coding sequence (Goedhart et al., 2012); provided by Joachim Goedhart) was amplified using primers mTU2_P2RP3_F and mTU2_P2RP3_wSTOP_R primers and recombined into pDONR-P2RP3 vector in a BP reaction to give mTURQUOISE2/pDONR-P2RP3. This plasmid was amplified using the INS_mTU2_P2RP3_F and INS_mTU2_P2RP3_wSTOP_R primers and the BB_mTU2_ P2RP3_F and BB_mTU2_ P2RP3_R primers. The PCR products were assembled by Gibson cloning (New England Biolabs) to give 2×mTurqoise2/pDONR-P2RP3. The UBQ10:H2B-mTurquoise2 transgene was introduced to Col-0 plants as described (Simon et al., 2014).

Genomic regions containing each of the TIR1/AFB genes were amplified using Phusion polymerase (New England Biolabs or ThermoFisher) from corresponding genomic clones (JAtY51F08, JAtY62P14, JAtY53F15, JAtY61O12, and JAtY52F19) except for AFB3 which was amplified from Col-0 genomic DNA. See Supplementary file 4 for primers used. The PCR products were cloned into pMiniT (New England Biolabs), and the stop codon was altered to create a NheI site using site-directed mutagenesis. An XbaI fragment containing either mCitrine (Griesbeck et al., 2001), mOrange2 (Shaner et al., 2008), or mCherry (Shaner et al., 2004) preceded by a short linker (either Arg-Gly₅-Ala or Arg-Gly₄-Ala) was ligated into the NheI sites. The genomic regions including the fluorescent protein genes were inserted in the MluI site of pMP535 (Prigge et al., 2005) as AscI fragments (AFB5) or as MluI-AscI fragments (others). To produce the sextuple-complementation construct, the TIR1-mOrange2 fragment was cloned into pMP535 as above, then AFB2-mCitrine was inserted into the re-created MluI site followed by AFB5-mCherry into its re-created MluI site. The constructs were introduced into the following strains by floral dip (Clough and Bent, 1998): tir1/+ afb5/+ afb1234 progeny (sextuple-complementation construct), tir1afb23 (TIR1-mCitrine, AFB3-mCitrine, and AFB3-mEGFP), tir1afb1245 (AFB2-mCitrine), afb45 (AFB4-mCitrine and AFB4-tdTomato), afb5-5 (AFB5-mCitrine, and afb1-3 (AFB1-mCitrine). Basta-resistant candidate lines were selected based on complementation of visible phenotypes (except for AFB1-mCitrine) then crossed to get them into the appropriate mutant backgrounds. Once in the sextuple-mutant background, the complementation transgene was maintained as a hemizygote by checking siliques for aberrant embryos or aborted seeds. The afb5-5 AFB5-mCitrine #9 and #19 lines were described previously (Prigge et al., 2016).

For confocal microscopy of the root meristem, five- to seven-day-old seedlings were stained in a 10 µg/ml aqueous solution of propidium iodide for one minute, rinsed in water, mounted with water, and viewed with either a Zeiss LSM 880 inverted microscope or a Zeiss LSM 710 inverted microscope. Embryos were fixed and stained with SCRI Renaissance 2200 (SR2200; Renaissance Chemicals, UK; Crawford et al., 2015). Briefly, using fine forceps and a 27-gauge needle as a scalpel, developing seeds were dissected from siliques and immediately immerged in fix solution (1 × PBS, 4% formaldehyde (Electron Microscopy Sciences, 15713), and 0.4% dimethyl sulfoxide) in a six-well plate with 100µ-mesh strainers. A vacuum was pulled and held three times for 12 min each time, before rinsing twice with 1 × PBS for 5 min. The embryos were transferred to SR2200 stain [3% sucrose, 4% diethylene glycol, 4% dimethyl sulfoxide and 1% SR2200 and stained overnight with vacuum pulled and released 3–4 times. Seeds were mounted (20% glycerol, 0.1 × PBS, 0.1% dimethyl sulfoxide, 0.1% SR2200, and 0.01% Triton X100) and the embryos were liberated by pressing on the coverslip. To detect mCitrine in the shoot apices, we removed stage 5 and older floral buds using fine forceps, fixed and rinsed (as with the embryos), soaked in ClearSee (Kurihara et al., 2015) for seven to ten days changing the solution every two to three days, and then stained with basic fuchsin (not shown) and Fluorescent Brightener 28 (Calcofluor White M2R) as described (Ursache et al., 2018). Confocal image channels were merged using ImageJ or FIJI (Schindelin et al., 2012; Schneider et al., 2012). Cleared embryos were viewed by mounting dissected ovules in a solution containing 2.5 g chloral hydrate dissolved in 1 ml 30% glycerol and viewed with a Nikon E600 microscope.

In order to infer the amounts of TIR1/AFB protein inside and outside the nucleus, 40 × magnification images of epidermal cells in the elongation zone of each TIR1/AFB-mCitrine lines were captured. Because the nuclei of AFB1-mCitrine-expressing cells are not apparent, the F₁ of a cross with a plant with a UBQ10:H2B-mTurquoise2 transgene was used to delineate the nucleus. Using FIJI (Schindelin et al., 2012), regions of interests including the entire cell (cell, based on propidium iodide staining), the nucleus (nuc, based on mCitrine or mTurquoise2 signal), and a cell-sized region outside the root (bg, background) were drawn using the freehand selections tool, and the area and mean gray values were measured for the mCitrine channel for each. The percent nuclear was calculated using the equation %_nuc = [Area_nuc × (Mean_nuc – Mean_bg)] ÷ [Area_cell × (Mean_cell – Mean_bg)].

The viable tir1afb lines were divided based on whether they contained the tir1 mutation, and the two batches were grown sequentially. The afb123 line included in the initial batch displayed a long-hypocotyl phenotype that may have been picked up after an earlier cross to the afb4-2 mutant, so a third batch was made up of alternative isolates for five lines whose pedigrees included a cross to afb4-2. Each batch included Col-0 and tir1-1. Seeds were surfaced sterilized, stratified in water for five days, spotted onto ½ Murashige and Skoog (MS) medium containing 1% sucrose, and incubated in a light chamber (22°C). Twelve five-day-old seedlings for each genotype were transferred to 120 mm square plates containing the same medium containing either 0, 20, or 100 nM IAA (batch a), 0, 100, or 500 nM IAA (batch b), or 0, 20, 100, or 500 nM IAA (batch c). Each plate received six seedlings from six genotypes spread out over two rows. Seedlings for each genotype were present on the top row of one plate and the lower row on a second plate placed in a different part of the growth chamber after marking the position of the root tips with a marker and scanning with Epson V600 flatbed scanners. The plates were scanned again after 72 hr (96 hr for batch c), and the growth was measured using imageJ. The plates containing 100 nM IAA were grown for a fourth day before the numbers of lateral roots protruding through the epidermis were counted using a dissecting microscope. Five seedlings from the no-IAA control plates were transferred to soil in 6 cm pots and grown an additional 34 days. The genotypes for two plants per line were confirmed by PCR. For each 42-day-old plant, the height from the rosette to the tip of the longest inflorescence and the maximum rosette diameter were measured, and the numbers of branches of at least 1 cm were counted. The IAA effects on root elongation data is presented as the percent relative to the growth without IAA ± the relative standard error of the ratio. For the gene effect analyses, the averages from each batch were normalized using measurements for Col-0 and tir1-1 plants that were included with each batch.

Seeds were sown on ½ MS medium containing 1% sucrose and 0.8% agar and stratified for 2–3 days at 4°C. Approximately fifteen 5-day-old seedlings were transferred to culture chambers (Lab-Tek, Chambers #1.0 Borosilicate Coverglass System, catalog number: 155361) containing the same agar medium supplemented with DMSO or IAA 10 nM (stock solution at 10 µM in DMSO). The transfer of seedlings was completed within 45–60 s. Images were acquired every 25 s for 20 min representing 50 images per root using Keyence microscope model BZ-X810 with 4 × lens.

Images obtained for one field were stacked and cropped to the region of interest (ROI). An auto threshold using the method ‘Default’ was applied. In addition, the ‘erode’, ‘despeckle’ and ‘Remove outliers’ (radius 10, threshold 50) functions were used to smooth the image and remove the remaining background. Each root tip was selected and the ‘Feret Distance’ within the ROI (which corresponds to the longest distance in an object) was determined for each root. Image processing was automated with an ImageJ macro, Supplementary file 8. For each time point the ‘Feret Distance’ root growth was calculated by subtracting the initial ‘Feret Distance’. The values obtained were used to generate graphs. For each genotype, the experiment was repeated three independent times.

To determine the effect of auxin on root growth throughout the experiment, the area under each curve of auxin-treated roots was determined and divided by the corresponding value for roots grown on DMSO condition to calculate the root growth response to IAA. A response value of 1 indicates that IAA had no effect on root growth. The effect of IAA on root growth was determined this way to account for differences in root growth between genotypes on DMSO.

Each sample was subjected to four different normality tests (Jarque-Bera, Lilliefors, Anderson-Darling and Shapiro-Wilk). Samples were considered as a Gaussian distribution when at least one test was significant (p=0.05). As a normal distribution was observed a one-way ANOVA coupled with a post hoc Tukey Honestly Significant Difference test was performed (p=0.05).

Measurements of etiolated hypocotyl elongation were carried out essentially as described previously (Fendrych et al., 2016; Li et al., 2018). Seeds were sterilized and stratified for four (set 1) or five (set 2) days before plating. After 6 hr of light treatment, the plates were wrapped in aluminum foil and sealed in a cardboard box for 66 hr. The plates were opened in a room lit only with an LED desk lamp with six layers of green cello film (Hygloss Products) filtering the light. Using a dissecting microscope with its light source filtered with six sheets of green cello film, the roots and cotyledons were excised using razor blades and the hypocotyls transferred to plates containing depletion medium (DM: 10 mM KCl, 1 mM MES pH 6, 1.5% phytagel) overlain with a piece of cellophane (PaperMart.com). After 30 to 80 min on DM, the hypocotyl segments were transferred to treatment plates (DM plus either 5 µM NAA or the equivalent amount of solvent (0.025% ethanol). Eight to sixteen hypocotyls were transferred for each genotype and treatment except for there being only five control-treated tir1afb23. Using Epson V600 flat-bed scanners, the plates were scanned at 1200 dpi 30–60 s after transfer then every ten minutes for three hours. The segments were measured using a FIJI macro that applied ‘Auto Threshold’ (Default), ‘Despeckle,’ ‘Remove Outliers’ (radius = 2 threshold = 50 which = Bright), then returned the ‘Feret Distance’ for each. For each segment at each time point, the Feret distance was subtracted from the initial Feret distance. The lengths were converted to µm using the conversion 21.16667 µm/pixel. In the second experiment, Col-0 hypocotyls were dissected first and a second batch was dissected after the other genotypes to test whether the length of time on DM affected the assay. Measurements for the two batches were only different at the 20 min time point (p<0.05 in two-tailed t-test).

In the experiments corresponding to Figure 7, six-day-old seedlings were positioned on four 120 mm square plates such that four seedlings of each genotype were in different positions in the four plates to reduce position effects. The plates were placed in the growth chamber vertically for an hour, scanned with an Epson V600 flat-bed scanners, then returned to the chamber vertically but rotated 90° from the original orientation. Plates were re-scanned every 30 min for 8 hr. For each root tip at each time point, the angle was measured in FIJI by drawing a line drawn from the medial point two-root-widths from the root tip to the root tip. These angles were corrected for scan-to-scan differences in plate orientation by measuring the angle of a horizontal line on the plate in each image. The mean changes in root-tip angle from that at time zero ± S.E.M. for each genotype at each timepoint was plotted. The experiments corresponding to Figure 7—figure supplement 1 were carried out in the same manner except that each plate contained a single genotype, and the seedlings were not repositioned onto different plates prior to rotation and scanning.