These authors contributed equally to this work.
Cellular efflux of auxin catalyzed by the Arabidopsis MDR/PGP transporter AtPGP1
Article first published online: 30 AUG 2005
The Plant Journal
Volume 44, Issue 2, pages 179–194, October 2005
How to Cite
Geisler, M., Blakeslee, J. J., Bouchard, R., Lee, O. R., Vincenzetti, V., Bandyopadhyay, A., Titapiwatanakun, B., Peer, W. A., Bailly, A., Richards, E. L., Ejendal, K. F. K., Smith, A. P., Baroux, C., Grossniklaus, U., Müller, A., Hrycyna, C. A., Dudler, R., Murphy, A. S. and Martinoia, E. (2005), Cellular efflux of auxin catalyzed by the Arabidopsis MDR/PGP transporter AtPGP1. The Plant Journal, 44: 179–194. doi: 10.1111/j.1365-313X.2005.02519.x
- Issue published online: 30 AUG 2005
- Article first published online: 30 AUG 2005
- Received 11 April 2005; revised 17 May 2005; accepted 10 June 2005.
- multi-drug resistance;
- ABC transporter
- Top of page
- Experimental procedures
- Supporting Information
Directional transport of the phytohormone auxin is required for the establishment and maintenance of plant polarity, but the underlying molecular mechanisms have not been fully elucidated. Plant homologs of human multiple drug resistance/P-glycoproteins (MDR/PGPs) have been implicated in auxin transport, as defects in MDR1 (AtPGP19) and AtPGP1 result in reductions of growth and auxin transport in Arabidopsis (atpgp1, atpgp19), maize (brachytic2) and sorghum (dwarf3). Here we examine the localization, activity, substrate specificity and inhibitor sensitivity of AtPGP1. AtPGP1 exhibits non-polar plasma membrane localization at the shoot and root apices, as well as polar localization above the root apex. Protoplasts from Arabidopsis pgp1 leaf mesophyll cells exhibit reduced efflux of natural and synthetic auxins with reduced sensitivity to auxin efflux inhibitors. Expression of AtPGP1 in yeast and in the standard mammalian expression system used to analyze human MDR-type proteins results in enhanced efflux of indole-3-acetic acid (IAA) and the synthetic auxin 1-naphthalene acetic acid (1-NAA), but not the inactive auxin 2-NAA. AtPGP1-mediated efflux is sensitive to auxin efflux and ABC transporter inhibitors. As is seen in planta, AtPGP1 also appears to mediate some efflux of IAA oxidative breakdown products associated with apical sites of high auxin accumulation. However, unlike what is seen in planta, some additional transport of the benzoic acid is observed in yeast and mammalian cells expressing AtPGP1, suggesting that other factors present in plant tissues confer enhanced auxin specificity to PGP-mediated transport.
- Top of page
- Experimental procedures
- Supporting Information
Transport of the plant auxin indole-3-acetic acid (IAA) is best described by a chemiosmotic model in which plasma membrane ATPases generate an H+ gradient between the neutral cytoplasm and the acidic extracellular space (Lomax et al., 1995). Cellular IAA uptake is mediated by lipophilic diffusion of IAAH augmented by tissue-specific gradient-driven H+ symport activity (Lomax et al., 1985; Swarup et al., 2004). The same gradient motivates carrier-mediated efflux of cytoplasmic anionic IAA−. The bias of auxin transport is attributed to highly regulated, polar-localized efflux complexes characterized by the PIN-FORMED (PIN) family of facilitator proteins (Friml and Palme, 2002). PINs have been shown to align with the auxin transport vector and to be necessary for normal polarized organ development and auxin movement (Benkova et al., 2003; Blilou et al., 2005; Galweiler et al., 1998; Palme and Galweiler, 1999). Treatment with auxin efflux inhibitors (AEIs) phenocopies some pin mutant phenotypes (Friml and Palme, 2002). Additionally, some members of the PIN family exhibit vectoral relocation during tropic growth, and function in the generation and maintenance of auxin sinks (Chen et al., 1998; Friml et al., 2002a,b; Muller et al., 1998).
Plant homologs of human multiple drug resistance/P-glycoproteins (MDR/PGPs) are numerous. The Arabidopsis PGP sub-family of ABC transporters is large, containing 21 members (five of which are highly homologous) (Jasinski et al., 2003; Martinoia et al., 2002). Structural characteristics of mammalian MDR/PGPs are well conserved in plant homologs, except in the predicted pore-facing helical domains thought to confer substrate specificity (Ambudkar et al., 2003). Analysis of expression patterns of the 21 members of the PGP subfamily, using both the AREX and AtGENEXPRESS microarray databases, shows that members of the PGP family exhibit distinct yet overlapping expression patterns (http://www.arexdb.org/index.jsp; Schmid et al., 2005).
PGPs have been implicated in auxin transport, as defects in MDR1 (AtPGP19) and AtPGP1 result in reduced growth and auxin transport of varying severity in Arabidopsis (atpgp1, atpgp19), maize (brachytic2/zmpgp1) and sorghum (dwarf3/sbpgp1) (Geisler et al., 2003; Multani et al., 2003; Noh et al., 2001). Further, auxin transport defects and dwarf phenotypes are more exaggerated in Arabidopsis double mutants, suggesting overlapping function (Noh et al., 2001). As gradient-driven anion efflux is sufficient to drive polar transport, it is not clear where plasma membrane-localized MDR/PGP ATP-binding cassette transporters fit into this scheme.
A mechanistic explanation of PGP function in auxin transport was suggested when, after fixation and detergent treatment, PIN1 was found to be mislocalized from the plasma membrane in xylem parenchyma cells of hypertropic Arabidopsis atpgp19 hypocotyls, but not in atpgp1 hypocotyls which do not exhibit altered tropic responses (Noh et al., 2003). These results suggest that PGPs might regulate transport by stabilizing plasma membrane efflux complexes, especially as PGPs are difficult to solubilize; are localized in detergent-resistant membrane microdomains (lipid rafts); bind the non-competitive AEI 1-N-naphthylphthalamic acid (NPA); and are components of membrane complexes that can be dissociated by NPA treatment (Geisler et al., 2003; Murphy et al., 2002; Noh et al., 2001). It is important to characterize the transport activity of PGPs in order to understand their functional role in these complexes.
Here we characterize the AtPGP1 protein (hereafter referred to as PGP1) by examining PGP1 localization, activity, substrate specificity and inhibition of activity. We confirm that PGP1 has an intermediate dwarf phenotype under short-day conditions, and characterize other auxin-related phenotypes. We show that PGP1 exhibits non-polar subcellular localization at the shoot and root apices that is consistent with PGP1 expression. We show that pgp1 mutant protoplasts exhibit reduced IAA transport, substrate specificity, and reduced sensitivity to AEIs. We show that PGP1 expressed in yeast and the standard mammalian cell expression system used to analyze human MDR-type proteins can mediate efflux of IAA and the synthetic auxin 1-NAA, but not the weak synthetic auxin 2-NAA. Further, PGP-mediated efflux is sensitive to auxin efflux and ABC-transporter inhibitors. PGP1 expression in yeast also results in increased resistance to toxic indolic IAA analogs. Expression of PGP1 in mammalian cells does not enhance the efflux of other classes of compounds that are common substrates for mammalian MDR-type transporters.
- Top of page
- Experimental procedures
- Supporting Information
pgp1 exhibits subtle auxin-related phenotypes
While the growth phenotypes of atpgp19 (hereafter referred to as pgp19) and the double mutant pgp1 pgp19 are clearly visible in both short- and long-day conditions (Geisler et al., 2003; Noh et al., 2001), those of pgp1 are not. Enhanced tropic bending observed in pgp19 hypocotyls is not observed in pgp1 hypocotyls (Noh et al., 2003), and gravitropic defects are difficult to quantify in pgp1 roots, largely because of greater variability in root gravitropic bending and root nutation compared with the wild type (Noh et al., 2003). Reduced hypocotyl growth reported in antisense PGP1 transformants (Sidler et al., 1998) was not observed in the pgp1 mutant (Noh et al., 2001) and, in our hands and under long-day conditions, shoot and hypocotyl growth of pgp1 mutants was difficult to distinguish from wild type (not shown). However, under shorter-day conditions, mature pgp1 exhibited an intermediate dwarf phenotype that is not as severe as pgp19 (Figure 1a) and is consistent with intermediate reduction in the transport of 3H-IAA from the shoot apex to the root–shoot transition zone previously observed in pgp1 (Geisler et al., 2003). However, as previous assays utilizing higher concentrations of 14C-IAA indicated slight increases in pgp1 auxin transport levels (Noh et al., 2001), free IAA levels in hypocotyls and whole roots of 5-day pgp1 and pgp19 seedlings were determined (Figure 1b) and were found to be consistent with more recently published transport data (Geisler et al., 2003).
Consistent with auxin levels observed in whole-root tissues, free IAA levels in pgp19 primary root tips were severely reduced (62 ± 18.3% of wild type). However, levels in pgp1 root tips were lower than expected (38 ± 29.7% of wild type). A small increase in IAA leakage from pgp1 (and not pgp19) root tips is also apparently a factor as, like the flavonoid-deficient mutant tt4 (Murphy et al., 2000; Peer et al., 2004), pgp1 root tips exhibited enhanced leakage of radiolabeled IAA into the support media in assays of shoot-to-root polar 3H-IAA transport (124 ± 9.4% of wild-type levels). Confirming these results, expression of the auxin reporter construct ProDR5:GUS was reduced in pgp1, pgp19 and pgp1 pgp19 compared with wild type, but was stronger in pgp19 than in pgp1 and pgp1 pgp19 (Figure 1c). Additionally, a stelar ProDR5:GUS signal observed in pgp19 was not visible in pgp1, consistent with a lesser accumulation of auxin in pgp1 root tips compared with pgp19. The reductions of ProDR5:GUS staining observed in pgp mutant root tips have been further confirmed in a recently published study showing reduced ProDR5:GUS expression in the root tips of independently generated pgp1 and pgp19 mutants (Lin and Wang, 2005).
IAA root basipetal transport in 5-day pgp1 seedlings
ProDR5:GUS results suggested that basipetal transport in pgp1 and pgp19 might be reduced at the root tip. Consistent with these results, export of radiolabeled IAA to the 2-mm segment proximal to the root tip in pgp1 and pgp19 was less than in the wild type (Figure 1d). Auxin transport to the next 2-mm segment was less than wild type in pgp19 but not pgp1, consistent with stronger PGP19 expression in non-apical tissues compared with PGP1 (Noh et al., 2001; Sidler et al., 1998) and greater apparent auxin retention in pgp19, indicated by ProDR5:GUS (Figure 1c). As PIN1 and PIN2 were not mislocalized in pgp1 root tips (Figure 2a–d), altered PIN localization cannot account for the altered auxin transport. Application of IAA ≥2 mm above root tips of both pgp mutants resulted in only marginal reductions in transport compared with wild type (approximately 5% in pgp19 and 14% in pgp1), suggesting limited PGP function in root basipetal transport in non-apical tissues. However, wild-type rates of basipetal auxin transport in these tissues are also much lower than apical transport rates, and decrease with distance from the root tip (data not shown). These results are similar to ProDR5:GUS activity and root basipetal transport reported in agr1-5/pin2 root tips (Shin et al., 2005).
PGP1 localization is non-polar at shoot and root apices
Auxin transport profiles of pgp1 and the strong expression of PGP1 observed in light-grown root and shoot apices (Noh et al., 2001; Sidler et al., 1998) focused our attention on PGP1 function in these regions. Unlike PGP19, which is highly expressed in upper hypocotyls and throughout the root in light-grown seedlings (Noh et al., 2001), PGP1 is expressed at lower levels in non-apical tissues in both light- and dark-grown seedlings (Geisler et al., 2003; Noh et al., 2001). ProPGP1:GUS enzyme and quantitative real-time PCR assays confirmed that PGP1 expression is strongest in root and shoot apices in dark-grown wild-type seedlings (Table 1). ProPGP1:GUS visualization also confirmed previous reports of PGP1 expression in lateral root primordia and apices (Sidler et al., 1998; Figure 3c), as well as previously unpublished weaker expression in cortical cells of the mature root and endodermal cells at the upper border of the distal elongation zone (Figure 3e). PGP1 expression in these tissues was further confirmed by examination of original in situ hybridization materials from Sidler et al. (1998) and microarray expression data in AREX (http://www.arexdb.org/index.jsp).
|Light||1.00 ± 0.73|
|Light + NPA||0.60 ± 1.66|
|Dark||14.89 ± 0.98*|
|Dark + NPA||3.70 ± 1.69|
|Wild type||1.0 ± 0.8|
|Wild type + IAA||8.5 ± 1.4*|
|Wild type||1.0 ± 0.9|
|Wild type + IAA||2.1 ± 0.9|
PGP1 proteins were immunolocalized utilizing a functional ProPGP1:PGP1-cmyc transformant (see Experimental procedures), and the localization patterns were compared with ProPGP1:GUS expression. PGP1 exhibits a strong non-polar localization in shoot and root apical cells as well as lateral root tips (Figure 3a,b,d). This suggests a role for PGP1 in non-directional auxin export from apical cells, and suggests that PGP function may be additive to, or synergistic with, PIN protein function.
Interestingly, in root tissues above the distal elongation zone, an apparent polar PGP1 localization was observed in mature cortical and endodermal cells at the upper boundary of the distal elongation zone (Figure 3f), suggesting that PGP1 may function in polar or reflux auxin movement (Blilou et al., 2005) in these tissues. In endodermal cells, the localization was always basal. In cortical cells, the localization was predominantly basal, although an apical localization without any obvious pattern was observed in some cortical cells flanking the stele. No reorientation of the basal signals was observed after microdeposition of IAA at the shoot or root tip or along the root surface (see Experimental procedures), suggesting a developmental basis for apical or basal localization. Transformation of pgp1 with ProPGP1:PGP1-cmyc complemented the mutant phenotype and restored wild-type auxin transport profiles (Figure 4c), and PGP1 protein localization was similar to that seen in transformed wild type (Figure 3g); PIN2 localization was not altered in transformants (Figure 3h). No signal was observed in wild-type immunolocalizations utilizing only primary or secondary antibodies under any detergent-solubilization conditions; no signal was observed in transformant immunolocalizations utilizing secondary antibodies only (Figure S1).
PGP1 expression is auxin responsive
As is the case for PGP19 (Noh et al., 2001), PGP1 expression appears to be auxin-responsive (Figure 3i,j; Table 1) and the PGP1 promoter contains auxin response element motifs (ARFAT, ASF-1 and NtBBF1). However, PGP1 expression in shoot tips did not expand spatially with auxin treatment (Figure 3i). PGP1 expression was also NPA-sensitive, and NPA treatment reversed increased PGP1 expression observed in dark-grown wild-type seedlings (Table 1).
PGP1 mediates cellular efflux in Arabidopsis protoplasts
In order to determine whether differences in auxin transport could be observed at the cellular level, efflux from protoplasts under conditions that minimize IAA catabolism was quantified. Protoplasts were isolated from leaf mesophyll cells of wild-type, pgp1, pgp19 and pgp1 pgp19 plants, but not from pin1, pin2, pin3 or pin4 mutants, as PIN1, PIN2, PIN3 and PIN4 expression is low in wild-type leaves (Figure 4a).
Wild-type protoplasts exhibited 3H-IAA efflux into the media (Figure 4b), and reductions in 3H-IAA efflux from pgp protoplasts compared well with transport reductions seen in whole plants: 72% in pgp1; 57% in pgp19; 49% in pgp1 pgp19 (Figure 4b). pgp1 mutants transformed with ProPGP1:PGP1-cmyc had efflux levels similar to wild type (Figure 4c). Vacuolar pH, relative protoplast volume and surface area, and chloroplast number per protoplast did not differ significantly between wild-type and pgp protoplasts, excluding indirect effects such as vacuolar trapping (Table 2). Consistent with ATP dependence of the process, auxin efflux was diminished in protoplasts isolated from plants kept in the dark for 24 h (ATP-depleted) (30% of light-grown wild type; Figure 4b).
|Vacuolar pH||Relative volume||Relative surface area||Number of chloroplasts|
|Wild type||6.40||1||1||37 ± 9|
|pgp1||6.31||0.976 ± 0.031||0.949 ± 0.015||40 ± 9|
|pgp19||6.19||0.998 ± 0.028||0.919 ± 0.020||36 ± 5|
|pgp1 pgp19||6.04||0.724 ± 0.053*||0.514 ± 0.058*||37 ± 9|
The synthetic auxin 1-NAA was exported by wild-type and pgp19 protoplasts, but not pgp1 or pgp1 pgp19 protoplasts (Figure 4b), suggesting that PGP1 transports this synthetic auxin better than PGP19. Substrate specificity in the protoplast system was investigated further using radiolabeled benzoic acid (14C-BA), a weak acid commonly used as a poorly transported control in plant assays. Negligible efflux of 14C-BA out of wild type, pgp1, pgp19, pgp1 pgp19, and ATP-depleted wild-type protoplasts was observed (Figure 4b), demonstrating that IAA efflux from protoplasts reflects the specific auxin transport observed in whole plants.
As the AEI NPA binds MDR/PGPs (Murphy et al., 2002; Noh et al., 2001), and NPA treatment disrupts membrane protein complexes containing PGPs (Geisler et al., 2003), IAA efflux from wild-type protoplasts would be expected to be NPA-sensitive, and pgp mutant protoplasts would be expected to exhibit diminished NPA sensitivity. However, an inactivating NPA amidase activity associated with the membrane aminopeptidase AtAPM1 (Katekar and Geissler, 1979, 1980; Katekar et al., 1981; Murphy and Taiz, 1999a,b; Murphy et al., 2002) is present in Arabidopsis leaves and is particularly active in leaf protoplasts (Murphy et al., 2002; Table 3). Treatment with NPA would be expected to increase initial 3H-IAA loading, but to be less effective in inhibiting 3H-IAA efflux. Treatment with NPA resulted in an 82% increase in 3H-IAA loading of wild-type protoplasts (Figure 4d), but only a 24% decrease in efflux. Consistent with even higher levels of NPA amidase activity in protoplasts derived from pgp1 and pgp19 (Table 3), measurements of NPA efflux inhibition from pgp protoplasts were highly variable.
|Time (min)||α-Naphthylamine formation (naphthylamine equivalents)|
|5||0.4 ± 0.91||3.2 ± 0.51|
|10||2.6 ± 2.21||6.8 ± 1.20|
|15||4.7 ± 2.34||7.1 ± 2.03|
However, when the non-hydrolysable NPA analog cyclopropyl propane dione (CPD) was substituted for NPA in wild-type protoplast assays, mean 3H-IAA efflux was reduced approximately 30% (Figure 4e). In contrast, CPD inhibited 3H-IAA transport from the tips of intact young Arabidopsis rosette leaves approximately 40% (not shown), suggesting that structural characteristics of intact tissues not present in protoplasts contribute to NPA/CPD sensitivity. Low levels of PIN expression in leaf protoplasts (Figure 4a) suggest that lower abundance of PIN proteins in mesophyll tissues may be involved in this reduction of AEI sensitivity. As expected, 3H-IAA efflux from CPD-treated protoplasts decreased approximately 12% in pgp1 and approximately 20% in pgp19 compared with the respective untreated protoplasts (Figure 4e). These results suggest that PGP19 is less sensitive than PGP1 to AEIs such as NPA and CPD, but also confirms that NPA/CPD inhibition requires factors that are altered in protoplasts.
The strong localization of PGP1 in apical tissues suggests that PGPs might also transport IAA breakdown products produced in these tissues (Kerk et al., 2000). Unlabeled, presumably partially degraded IAA is often included in whole-plant radiolabeled auxin transport assays as it enhances measurable transport, possibly by enhancing uptake (Rashotte et al., 2000). Protoplast assays were repeated using either >99% radiolabeled IAA or a 1:1 mixture of IAA and IAA oxidative breakdown products (primarily oxindoleacetic acid and indoleacetaldehyde, as determined by LC/MS and GC/MS). Increased loading and efflux was observed with the mixture (Figure 4f), but relative efflux from wild-type and pgp mutant protoplasts at 10 min was similar with both treatments. These results suggest that, in leaf cells, uptake and efflux of some IAA metabolites may be mediated by a PGP-dependent mechanism. However, it is not clear how far the specificity of IAA efflux might be enhanced in tissues where PINs are strongly expressed.
Analysis of PGP1 and auxin efflux in yeast
Heterologous expression of PIN2 in yeast was previously used to demonstrate a positive impact on net auxin efflux (Chen et al., 1998). Although PGP19 was found to be mistargeted when expressed in a Saccharomyces cereviseae deletion strain (JK93da) with reduced ABC transporter activity (Noh et al., 2001), immunohistochemical analyses indicated that PGP1 expressed in JK93da strain was localized in the plasma membrane (Figure 5a). Yeast expressing PGP1 exhibited time-dependent loading and efflux of radiolabeled IAA, 1-NAA and BA (Figure 5b). 3H-IAA efflux was temperature- and glucose-dependent (Figure S2a,b); membrane permeability was unaltered (Figure S2c); and the effluent species was determined by GC-MS to be 3H-IAA (Figure 5d). Efflux of 3H-IAA was seven times greater in PGP1-transformed yeast than in vector controls, and efflux of 3H-1-NAA was also slightly increased (Figure 5b). However, although no efflux of the weak synthetic auxin 2-NAA was seen (Figure 5c), a lack of specificity was indicated when efflux of 14C-BA was seen to be almost equal to that of 3H-IAA (Figure 5b).
Consistent with PGP1 mediation of auxin efflux, 3H-IAA efflux in PGP1-transformed yeast was significantly reduced by treatment with NPA, the mammalian MDR/PGP inhibitors cyclosporin A and verapamil, and the inhibitor of both auxin transport and mammalian MDR/PGPs, quercetin (Figure 5e). Further, expression of PGP1 in the hypersensitive gef1 yeast mutant exhibited increased resistance to the cytotoxic IAA analog 5-fluoroindole which could be reversed by both AEIs and ABC transport inhibitors (M.G., unpublished data). Additionally, yeast yap1-1 mutants, which are hypersensitive to IAA due to increased expression of AUX1-like permeases (Prusty et al., 2004), are rescued by AtPGP1 but not by mouse MDR3 (Raymond et al., 1992) when grown on IAA (Figure 5f).
Analysis of PGP1-mediated auxin efflux in mammalian cell lines
Vaccinia virus/T7 RNA polymerase expression in human HeLa cells has become a standard system for assaying mammalian PGPs, as it provides efficient heterologous expression of functional eukaryotic plasma membrane proteins and suppression of host-protein synthesis (Elroy-Stein and Moss, 1991; Hrycyna et al., 1998; Moss, 1991). Protocols to confirm protein functionality and to assay net accumulation of radiolabeled and fluorescent substrates are well established (Hrycyna et al., 1998). These protocols were modified to allow loading of 3H-IAA into HeLa cells while maintaining cell viability and substrate integrity.
HeLa cells expressing PGP1 exhibited significant 3H-IAA efflux compared with empty vector controls (Figure 6a). Although loading of 3H-1-NAA was less than that seen with 3H-IAA, cells expressing PGP1 exhibited a small net efflux of 3H-1-NAA (Figure 6a). Cells expressing PGP1 exhibited significant 3H-IAA efflux compared with cells transformed with empty vector alone over a sixfold concentration range (Figure 6a,b). No net efflux of 2-NAA could be detected in cells expressing PGP1 (Figure 6c). However, as seen when PGP1 was expressed in yeast, PGP1-mediated net 3H-BA efflux was also observed when 3H-BA was supplied at higher concentrations (Figure 6b). Background efflux of 3H-BA in empty vector controls was higher than with 3H-IAA, presumably due to endogenous mammalian benzoate/monocarboxylic acid transport activity (Yan and Taylor, 2002). Expression of PGP1 in HeLa cells resulted in more auxin substrate specificity than when PGP1 was expressed in yeast, as PGP1-mediated net efflux of 3H-BA decreased to zero when 3H-BA concentration was reduced to 10 nm, while 3H-IAA efflux was unaffected (Figure 6b). Competition assays utilizing 10× more unlabeled BA than 3H-IAA resulted in substantial inhibition of background 3H-IAA efflux (data not shown), and 3H-IAA efflux was reduced 22 ± 12.5% less in cells expressing PGP1 than in empty vector controls. This suggests that background IAA efflux in HeLa cells is mediated by a monocarboxylic acid transporter, and that PGP1 is a functional IAA transporter when expressed in HeLa cells. Supporting this conclusion, the monocarboxylate transport inhibitor cardio green reduced background but not PGP1-mediated efflux (not shown).
All the assays were conducted in a buffer solution containing 1–2 mm citrate (see Experimental procedures), indicating that organic acids such as citrate and malate are not substrates for PGP1-mediated transport. Further, no efflux of mammalian MDR/PGP hydrophobic substrates (rhodamine 123, daunomycin, BODIPY-vinblastine) was seen in cells expressing PGP1 (Figure S3a–d). These results indicate that PGP1-mediated efflux is specific for active auxins and, to a lesser extent, a subset of aromatic carboxylic acids.
PGP1-mediated efflux of IAA oxidative degradation products in HeLa cells
As protoplast assays suggested that PGP1 may also be involved in efflux of IAA oxidative degradation products (Figure 4f), HeLa cells expressing PGP1 were incubated with a 1:1 mixture of 3H-IAA and oxidative IAA breakdown products. Consistent with protoplast results, incubation of cells with 10 nm (indole equivalents) of the mixture resulted in net loading in transfected cells to levels twice those observed when cells were loaded with 63 nm fresh IAA. PGP1-mediated net 3H-IAA efflux was also twice as high with this treatment (Figure 6d), but, NPA sensitivity observed at lower IAA concentrations was retained. When fresh 3H-IAA was mixed with an equal amount (indole equivalents) of unlabeled oxidative breakdown products, net efflux was reduced (Figure 6d), suggesting competitive inhibition of IAA efflux. These data support a role for PGP1 in the transport of auxin breakdown products in planta.
Inhibitor studies of PGP1-mediated efflux in HeLA cells
The effects of inhibitor treatments on PGP1-mediated 3H-IAA efflux were normalized to inhibitor-treated empty vector controls. PGP1-mediated 3H-IAA net efflux was inhibited by treatment with 10 μm NPA (Figure 6e), and efflux was more NPA-sensitive at lower IAA concentrations (Figure 6e,f). Both mammalian and Arabidopsis PGPs have been shown to bind the aglycone flavonoid quercetin (Ferte et al., 1999; Murphy et al., 2002) which decreases mammalian PGP activity and negatively regulates auxin transport in plant tissues where PGPs are expressed (Brown et al., 2001; Ferte et al., 1999; Peer et al., 2001, 2004). Treatment with flavonoids and mammalian MDR/PGP inhibitors has previously been shown to reverse efflux of human MDR substrates to the point of net retention in mammalian cells (Zhang and Morris, 2003). Net 3H-IAA efflux by PGP1 was reversed when cells were treated with 200 nm quercetin (Figure 6e,f). Treatment with the MDR/PGP inhibitors verapamil and cyclosporin A increased net retention in cells expressing PGP1 (Figure 6e,f), suggesting an additive cytotoxicity of these compounds in transfected cells. These data support the hypothesis that PGP1-mediated IAA transport can be regulated by endogenous flavonoids, as well as NPA. Apparently due to the background activities observed, PGP-mediated efflux exhibited reduced sensitivity to quercetin and standard MDR/PGP inhibitors at lower IAA concentrations (Figure 6e).
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- Experimental procedures
- Supporting Information
Taken together, these results indicate that, in Arabidopsis, PGP1 can mediate cellular efflux of IAA and some IAA metabolites. Further, protoplast and whole-plant transport assays suggest that PGPs are capable of mediating auxin transport in planta, although other transport proteins may also function in auxin efflux. As PGP1-mediated auxin efflux in heterologous systems exhibits decreased specificity and AEI sensitivity, other factors present in planta are probably required for the high degree of specificity seen in polar auxin transport. Non-polar localization of PGP1 at root and shoot apices suggests that interactions with other proteins are also required to confer directionality to auxin transport in those tissues. In these tissues, where rediffusion of IAA exported from small cells would be expected to impair gradient-driven transport, additional energy-dependent IAA efflux might be required.
PGP1 localization and inhibition by quercetin are also consistent with its proposed interaction with endogenous flavonoids (Murphy et al., 2000; Peer et al., 2004). Strong expression of PGP1 in apical tissues, localized reduction of IAA transport in the root tips of pgp1 mutants, and PGP-mediated transport of IAA breakdown products suggest that PGP1 functions in regions exposed to high auxin concentrations. Further, the non-polar localization of PGP1 in root apices suggests that, where PIN–PGP interactions might take place, PIN proteins may confer an accelerated vectoral component to PGP-mediated transport. Specific interactions of the many PIN and PGP family members in discrete tissues would be predicted to provide tight control of transport mechanisms. To elucidate the role of PGP family members in auxin transport, further developmental and cell biological studies are required.
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- Experimental procedures
- Supporting Information
Plant growth conditions
Expression and localization analysis
To construct the PGP1 (At2g36910) cmyc tag line (ProPGP1:PGP1-cmyc), a synthetic double-stranded oligonucleotide (forward primer: 5′-ctagagcagaagcttatctccgaggaggacct ctagtgagct; reverse primer: 5′-cactagtaggtcctcctcggagataagcttctgct) encoding the 9E10 c-myc epitope was introduced into SpeI and SacI sites of construct p4kbPstI (Sidler et al., 1998), leading to p4kbMyc2. The cmyc epitope is thereby inserted at amino acid residue Leu14. In comparison with the original sequence, a base pair was changed (bold letter) leading to introduction of a diagnostic HindIII site (underlined) without changing the codon bias. The 5′SpeI site of the oligonucleotide (italic) was designed in such a way that it was destroyed upon ligation, allowing us to use the downstream SpeI site (italic) for further cloning. To add the 3′-terminal part of the PGP1 gene, a 6.7-kb SpeI fragment was cut out from plasmid pOE and inserted in the correct orientation into p4kbMyc2 predigested with SpeI, leading to pBSKMdrMyc2. From this a 9-kb SalI fragment encompassing PGP1 was subcloned into the SalI site of binary plant transformation vector pBI101 (Clontech, Palo Alto, CA, USA). The final construct, PGP1cmyc2b (ProPGP1:PGP1-cmyc), containing the full-length genomic fragment of PGP1 (including the cmyc tag at base pair 980) flanked by the PGP1 promoter and terminator sequences, was introduced into Arabidopsis thaliana (ecotype Columbia) by Agrobacteriuhm tumefaciens-mediated vacuum infiltration. Homozygous plants were selected on 0.5× MS medium containing 50 μg ml−1 kanamycin, and insertion was verified by Southern analysis.
For complementation, pgp1 mutant plants (Noh et al., 2001) were transformed with the binary vector PGP1cmyc2b (ProPGP1:PGP1-cmyc) via vacuum infiltration, and homozygous lines were selected on 0.5× Murashige and Skoog (MS) medium containing kanamycin.
Arabidopsis seedlings were grown on 1% phytagar plates (0.5× MS basal salts, pH 4.85) under white light (photon flux rate 100 μmol m−2 sec−1, 14 h light, 22°C) and 5- and 9-day seedlings were prepared for immunolocalization as described by Friml et al. (2003) with the following modifications: in some cases, in place of 2% driselase, 0.5 pectolyase was used for 5-day seedlings and 0.8% pectolyase for 9-day roots. Seedlings were incubated overnight (37°C) with mouse anti-cmyc antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA) at 1:250 dilution, for 3 h at 37°C with rabbit anti-mouse-FITC (whole IgG, Sigma F7506, St Louis, MO, USA) and for 3 h at 37°C with goat anti-rabbit-Alexa 488 at 1:250 in 3% BSA/MTSB, respectively. Immunofluorescence analysis was done using a confocal laser scanning microscope (Nikon, Melville, NY, USA, Eclipse 800) equipped with an argon laser (488 nm) (Bio-Rad). Images were captured with a SPOT camera and processed using Adobe photoshop 5.0.
For histochemical GUS staining, 10-day light-grown ProPGP1:GUS transgenic seedlings were incubated in 10 mm HEPES (pH 5.2), 500 mm sorbitol containing 2 μm benzoic acid or IAA for 3 h. For all comparisons between treatments, identical staining conditions were used. For promoter gene quantification, ProPGP1:GUS transgenic seedlings were transferred for 3 h onto 0.5× MS plates supplemented with 1 μm IAA. Seedlings were washed in MS and separated manually into root and shoot, and GUS activity was assayed using 4-β-4-methylumbelliferone-glucuronide as substrate. For each sample, segments of 40 seedlings were pooled and analyzed.
Protoplast and cell culture efflux experiments
Arabidopsis mesophyll protoplasts were prepared from rosette leaves of plants grown on soil under white light (100 μmol m−2 sec−1, 8 h light/16 h dark, 22°C). Intact protoplasts were isolated as described (Geisler et al., 2003) and loaded by incubation with 1 μl ml−13H-IAA (specific activity 20 Ci mmol−1, American Radiolabeled Chemicals, St Louis, MO, USA), 7-14C-benzoic acid (53 mCi mmol−1, Moravek Biochemicals, Brea, CA, USA) or 4-3H-1-naphthalene acetic acid (25 Ci mmol−1, American Radiolabeled Chemicals) on ice. External radioactivity was removed by separating protoplasts using a 50–30–5% percoll gradient. Samples were incubated at 25°C and efflux halted by silicon oil centrifugation. Retained and effluxed radioactivity was determined by scintillation counting of protoplast pellets and aqueous phases. For inhibitor studies, protoplasts were isolated and assayed in the presence of 10 μm of indicated inhibitors or the solvent alone.
Efflux experiments were performed with three to five independent protoplast preparations with four replicas for each time point. Protoplast volumes were determined by the addition of 0.05 μCi 3H2O in separate assays; protoplast surfaces were calculated by measuring protoplast diameters. Vacuolar pH was determined directly from homogenated whole leaves following centrifugation.
For inhibitor studies, 50 ml Arabidopsis cell suspension culture (May and Leaver, 1993) was grown in MS basal medium supplemented with 3% sucrose (w/v), 0.5 mg l−1 naphthalene acetic acid, and 0.05 mg l−1 kinetin in the presence of 10 μm of indicated inhibitors or the solvent alone for 12 h. Prior to measurements, the culture was centrifuged for 10 min at 115 g and 4°C, washed with sterile water and resuspended in 10 ml MCP (500 mm sorbitol, 1 mm CaCl2, 10 mm MES, pH 5.6). Then 1 μl ml−13H-IAA (specific activity 20 Ci mmol−1, American Radiolabeled Chemicals) was added, and four 500 μl aliquots were collected after 0 and 10 min incubation at 25°C and filtered on Millipore Durapore 0.22 μm GV filters. Cylcopropyl propane dione (CPD) was obtained from Caisson Labs (Rexburg, ID, USA).
Transcript detection by RT-PCR
Total RNA from Arabidopsis wild-type protoplasts was prepared, and DNaseI (Qiagen, Valencia, CA, USA) treatment was performed with column-bound RNA. Oligo dT-primed cDNA from 1 μg total RNA was synthesized using the reverse transcription system (Promega, Madison, WI, USA). Transcripts specific for PGP1 (At2g36910), PIN1 (At1g73590), PIN2 (At5g57090), PIN3 (At1g70940), PIN4 (At2g01420) and 40S ribosomal protein S16 (At2g09990) were detected by conventional PCR for 30 and 35 cycles at 52°C annealing temperature. Intron-spanning PCR primers were: S16-S 5′-ggcgactcaaccagctactga; S16-AS 5′-cggtaactctt tggtaacga; PGP1-S 5′-gtccctcaagagccgtgcttg; PGP1-AS 5′-ccatcatcgatgacagcgatc, PIN1-S 5′-tggagctcaagtgcttcgccg; PIN1-AS 5′-gagaagagttatgggcaacgc; PIN2-S 5′-cacggggtcaacgagtggagc; PIN2-AS 5′-ctgagaatatcaggatggacg; PIN3-S 5′-tgatgacaaggctgatactg; PIN3-AS 5′-gtaaatcaccacgtaatccg; PIN4-S 5′-acaacgccgttaaatatgga; PIN4-AS 5′-agaccccattttattcagcc. Equal volumes of PCR products were separated on 2.5% agarose gels. Negative controls in the absence of enzyme in the reverse transcriptase reaction yielded no products.
RNA isolation and quantitative RT–PCR analysis
RNA isolation and quantitative RT–PCR analysis were carried out as described previously, with slight modification (Blakeslee et al., 2004), using the ABI Prism 7000 Sequence Detection System (Applied Biosystems, Foster City, CA, USA) in mixtures of 10 μl TaqMan Universal PCR Master Mix containing AmpliTaq Gold DNA polymerase, AmpErase uracil-N-glycosylase (UNG), deoxynucleoside triphosphates with dUTP, a passive reference dye, optimized buffer components (Applied Biosystems), 500 nm each primer and optimum amount of template DNA in a total volume of 20 μl. β-tubulin (At5g12250) forward and reverse primers were 5′-ttcccgggtcagctcaac and 5′-ggagacgagggaaaggaatga, respectively. Primers used for the AtPGP1 transcript were 5′-tctggcgactagctaaaatgaactc and 5′-ccacaaatgacagagcctactga. The AtPGP19 forward primer sequence was 5′-ggaagtttgaggaatctgagctattc, and the reverse primer sequence was 5′-tcggtcagtctctgcatttga. Transcripts specific for β-tubulin, AtPGP1 and AtPGP19 were detected by PCR; activation of AmpErase UNG (2 min, 50°C) and Taq polymerase (10 min, 95°C), 40 cycles of denaturation (15 sec, 95°C) and elongation (1 min, 60°C).
Analysis of IAA contents and transport
pgp1 mutants expressing the maximal auxin-inducible reporter ProDR5:GUS (Ulmasov et al., 1997) were generated via crossing with wild-type ProDR5:GUS plants. Seedlings were grown for 5 days as described above and stained for GUS expression (Ulmasov et al., 1997).
For endogenous free auxin quantification, seedlings were grown as previously described (Geisler et al., 2003). Nine days after planting seedlings were harvested, the cotyledons excised, and seedlings cut in half at the root–shoot transition zone. Roots and shoots were collected in lots of 500, and free auxin was quantified by GC-MS as described previously (Chen et al., 1988). Data presented are the averages of three lots of 500 seedlings. Auxin quantifications were confirmed by GC-MS after pentafluorobenzyl derivatization (Prinsen et al., 2000).
For additional free auxin quantifications, hypocotyl and root segments of 30–50 seedlings were collected and pooled. Samples were extracted and analyzed by GC-MS. Calculation of isotopic dilution factors was based on the addition of 100 pmol 2H2-IAA to each sample. In some cases, roots of 40 seedlings were divided manually into 2-, 8- and 10-mm segments from the root tip, and analyzed as described above.
Auxin transport assays on intact, light-grown Arabidopsis seedlings treated with a 0.1 μl microdroplet of 1 μm auxin at the root apical meristem, using techniques described by Geisler et al. (2003), and root segments of 2 mm, were collected 2, 4 and 6 mm from the root tip.
Triplicate samples were pooled and extracted in methanol/2% HCl with shaking at 4°C for 30 min. An equal volume of diethylether/hexane (1:1) was added, samples were shaken vigorously, and the upper phase was collected. This was repeated twice. Aminopropylsilyl solid phase extraction columns (Alltech, State College, PA, USA), were preconditioned with hexane before sample extracts were added. Eluate was collected from the application of three bed volumes of diethylether/2% formic acid to the column. Samples were dried prior to resuspension in methanol/1% phosphoric acid. Samples were analyzed by LC-MS using a Waters C18 MS column on a Q-TOF Micro equipped with a collision cell and phosphoric acid lock spray for internal calibration (Waters, Bedford, MA, USA), using naphthalene as an internal standard.
Auxin metabolite characterization
IAA oxidative breakdown products were determined by LC-MS and GC-MS (Ljung et al., 2002, 2005).
The PGP1 cDNA was cut out from plasmid pUCcMDR (Windsor et al., 2003) with StuI and HincII and cloned into the Klenow filled NotI site of yeast shuttle vector pNEV (pNEV-PGP1). pNEV and pNEV-PGP1 were introduced into S. cerevisiae strain JK93da or yap1-1 (Prusty et al., 2004), and single colonies were grown in synthetic minimal medium without uracil (SD-URA), supplemented with 2% glucose. For detoxification assays, transformants grown in SD-URA to an OD600 of approximately 0.8 were washed and diluted to OD600 = 1 in water. Cells were 10-fold diluted five times, and 5 μl of each dilution were spotted onto minimal media plates supplemented with 10 μm IAA. Growth at 30°C was assessed after 3–5 days. Assays were performed with three independent transformants.
For loading experiments, cells were grown to OD600 = 1, washed and incubated at 30°C with combinations of 1 μl ml−1 5-3H-IAA (specific activity 20 Ci mmol−1, American Radiolabeled Chemicals), 7-14C-BA (53 mCi mmol−1, Moravek Biochemicals), 4-3H-1-NAA (25 Ci mmol−1, American Radiolabeled Chemicals) or 1 μm 2-NAA (Sigma) in SD-URA (pH 4.5). Aliquots of 0.5 ml were taken after 0, 5, 10, 30, 60 and 120 min incubation at 30°C. For efflux experiments, cells were loaded for 10 min on ice, washed twice with cold water, and resuspended in 15 ml SD (pH 4.5). Then 0.5-ml aliquots were taken after 0, 2.5, 5, 7.5 10, 15 and 20 min incubation at 30°C.
For assaying permeabilities, yeast was grown to logarithmic phase (OD600 = 1.25), washed with water followed by fluorescein diacetate (FDA) buffer, and resuspended in 5 ml FDA buffer [50 mm HEPES-NaOH (pH 7.0), 5 mm 2-deoxy-d-glucose]. Then 10 μl 5 mm FDA was added to a 990 μl cell suspension, and aliquots were measured at 485 nm excitation and 535 nm emission. To determine the temperature dependency of efflux, loaded yeast cells were washed, divided into two aliquots and incubated at 4 and 30°C, respectively. For assaying ATP dependency of transport, yeast were grown to logarithmic phase, washed with water and FDA buffer, resuspended in 50 ml FDA buffer, and incubated for 1 h at 30°C. Cells were harvested, washed and resuspended in 10.8 ml 20 mm sodium citrate (pH 4.5). After temperature equilibration at 30°C, 1 μl ml−1 5-3H-IAA was added, and resuspensions were divided into two aliquots followed by addition of 0.6 ml 20% glucose to one aliquot. For inhibitor studies, yeasts were grown for 12 h prior to measurement, and assayed in the presence of 10 μm of indicated inhibitors or the solvent alone.
All aliquots were filtered on Whatman GF/C filters and washed three times with cold water, and retained radioactivity was quantified by scintillation counting. All transport experiments were performed three to five times with independent transformants, with four replicas each.
To verify the identity of effluxed IAA, yeast cells transformed with pNEV and pNEV-PGP1 were loaded with 10 μm2H5-IAA and 1 μm13C6-IAA in 20 mm sodium citrate (pH 4.5). Effluxed IAA was extracted by ethyl acetate and analyzed by MS-MS.
Yeast PGP1 expression and immunolocalization
Yeast cells transformed with pNEV and pNEV-PGP1 were grown to mid-log phase and microsomes were separated via 7.5% PAGE. Western blots were immunoprobed using anti-PGP1 antibody (Sidler et al., 1998). Yeast immunolocalization was performed using standard protocols. Fixed yeast cells were incubated overnight at 30°C with rabbit anti-PGP1 antibody (Sidler et al., 1998) and for 3 h with goat anti-rabbit-Alexa 488 at 1:200 in 3% BSA. Stained cells were incubated in mounting media containing 4′,6-diamidino-2-phenylindole (DAPI), and immunofluorescence analysis was done using a confocal laser scanning microscope (Leica, DMIRE2) equipped with argon (488 nm) and UV laser (410 nm). Fluorescence, differential interference contrast and DAPI images were processed using Adobe photoshop 7.0. Vector controls showed no detectable fluorescence with anti-PGP1 antibody.
HeLa cell assays
Radiolabeled substrate accumulation assay. PGP1 (At2g36910) was expressed in mammalian HeLa cells using a vaccinia virus co-transfection system providing several advantages over other heterologous expression systems, including proper glycosylation and suppression of host-protein synthesis following vaccinia infection (Elroy-Stein and Moss, 1991). The transient vaccinia expression system was used because stable cell lines develop mutations and express other endogenous drug-resistance mechanisms. Full-length, hemagglutinin (HA)-tagged PGP1 was cloned into the multiple cloning site of the pTM1 vector (Hrycyna et al., 1998). For pTM1-PGP1, a PGP1 PCR fragment containing XmaI/BamHI restriction sites was generated using the following primers: PGP1-S 5′-tcc ccc cgg ggc atg gat aat gac ggt and PGP1-AS 5′-cgc gga tcc agc gta atc tgg tac gtc gta agc atc atc ttc ctt aac. Assays for accumulation of radiolabeled substrates were performed according to the method described by Hrycyna et al. (1998), with the following modifications: cells were transfected in six-well plates with 2 μg DNA (pTM1 control vector, PGP1) per well. For radiolabeled substrate accumulation assays, gradient conditions were developed wherein radiolabeled auxin was passively accumulated by empty vector control HeLa cells without induction of cellular damage. Confluent cells were transfected in six-well plates, and 16–24 h after transfection cells were washed with 3 ml pre-warmed Dulbecco's modified Eagle's medium, 5% fetal bovine serum. Each transfection utilized 600 000–1 000 000 cells, and equal loading of wells was verified by sampled cell counts. Cell counts were determined by Coulter counting and microscopic visualization (percentage confluence). Cells were then incubated with 2 ml PBS citrate buffer pH 5.5, 5% calf sera containing either 10 or 62.5 nm of the following radiolabeled substrates: 3H-IAA (specific activity 26 Ci mmol−1, Amersham Biosciences, Piscataway, NJ, USA); 3H-benzoic acid (specific activity 20 Ci mmol−1, American Radiolabeled Chemicals); or 3H-1-NAA (specific activity 20 Ci mmol−1, American Radiolabeled Chemicals). Possibly due to buffer compatibility issues, it was difficult to maintain solubilization of 1-NAA in loading assays. For radiolabeled auxin degradation product assays, cells were loaded with 10 nm radiolabeled IAA breakdown products (specific activity 25 Ci mmol−1, American Radiolabeled Chemicals). Cells were incubated with radiolabeled substrates for 40 min at 37°C, 5% CO2. For inhibitor studies, cells were incubated with radiolabeled IAA in the presence of 10 μm NPA, 200 nm quercetin, 1 μm cyclosporin A or 5 μm verapamil. After incubation, cells were washed three times with 3 ml ice-cold PBS, removed from the wells by trypsinization, and added to 18 ml scintillation fluid. Samples were counted in a Perkin-Elmer scintillation counter. Components of the radiolabeled auxin breakdown product mixture were determined and quantified via LC-MS. For cold 2-NAA retention studies, cells were incubated with 62.59 nm cold 2-NAA, harvested and extracted. 2-NAA was quantified using LC-MS. As with 1-NAA, it proved difficult to keep 2-NAA solubilized for cell loading in the buffer system used. Fluorescent substrate accumulation assays were performed in the HeLa cell system as previously described (Hrycyna et al., 1998).
Data points were normalized to the average empty control vector value of 2851.885 DPM/500 000 cells for auxin treatments. Cell viability after treatment was confirmed visually and via cell counting.
Western blotting of HeLa cells
HeLa cells expressing PGP1 were harvested using a rubber policeman. Proteins were isolated from cells as described previously (Hrycyna et al., 1998). Proteins were separated via 7% SDS-PAGE, then transferred onto nylon membrane at 10 V, in an ice bath, overnight. After blocking with 5% non-fat dried milk in 1× PBS with 0.1% Tween 20, the nylon membrane was incubated with primary antibody for 2 h at 4°C (anti-HA, Sigma). Following washing, membranes were incubated with horseradish peroxidase-conjugated secondary antibody, and developed using the ECL chemiluminescence system according to the manufacturer's protocol (Amersham Pharmacia Biotech, Piscataway, NJ, USA).
Data were analyzed using prism 4.0b (Graphpad Software, San Diego, CA, USA). Statistical analysis was performed using spss 11.0 (SPSS Inc., Chicago, IL, USA) and sigmastat (Systat, Point Richmond, CA, USA).
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We thank V. Croy, C. Gaillard, C. Ringli and A. Hopf for technical assistance, S. Plaza for help with statistical analysis, and I. Baxter for microarray data. This publication is dedicated to M.D.R. Louma (MG) and to the memory of Kenneth Thimann (ASM and WAP). This manuscript was written by ASM, MG, JJB, WAP and EM. Experiments were conceptualized by MG, JJB, WAP, ASM and EM. Auxin transport assays – whole plant: JJB; protoplast: MG, RB, VV, AB, JJB and ASM; yeast: MG, RB, JJB and ASM; HeLa cells: JJB. Implementation of HeLa cell system for auxin transport studies: JJB, ASM, ORL, KFKE and CAH. Immunolocalizations – yeast: JJB, MG and VV; Arabidopsis PIN and PGP-cmyc: AB, BT, CB and UG. Auxin determinations – ASM, AM and ELR. Cloning of PGP1 into the pTM1 expression vector by ORL. Real-time PCR analysis of PGP1 expression in HeLa cells and whole plants by ORL and WAP. DR5::GUS/PGP reporter constructs prepared by AS and ASM; DR5::GUS assays by JJB and WAP; PGP1::GUS reporter constructs created by RD; PGP1::GUS assays by MG; PGP1-cmyc transformants created by MG. Funding: National Science Foundation (US) to ASM, National Science Foundation (SWISS) to EM.
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Figure S1. Anti-c-myc antibody controls. A. No immunolocalization signal was observed in wild type root tips with the anti-c-myc antibody. B. No immunolocalization signal was observed in wild type mature roots with the anti-c-myc antibody. Bar, 125 μm.
Figure S2. PGP1 expressed in yeast cells with reduced ABC transporter activity. As might be expected for an energized transport mechanism, PGP1-mediated IAA export in yeast was both temperature- and glucose-dependent (Fig. S2A, B). Yeast cell permeability was similar in cells transformed with either empty vector or PGP1 (Fig. S2C). A. IAA efflux in yeast is temperature-dependent. Initial loading was at the same magnitude (+/-10%). Presented data represent relative mean activities plus standard deviations. B. IAA efflux in yeast is glucose-dependent. Initial loading was at the same magnitude (+/-10%). Presented data represent relative mean activities plus standard deviations. C. Yeast transformed with pNEV-PGP1 (PGP1) or empty plasmid pNEV (control) have similar permeabilities. Yeast cells were incubated with the dye FDA and internal fluorescence was measured. Presented data represent relative mean activities of three to five individual measurements of four samples plus standard deviations.
Figure S3. PGP1 expressed in HeLa cells did not transport HsMDR substrates. A-D. HeLa cells expressing PGP1 did not transport common mammalian MDR substrates. Grey peak represents cells transformed with pTM1 control vector (negative control). Solid line, Homo sapiens MDR1 (positive control); large dashes, PGP1. Shifting of observed peaks to the left indicates transport of the fluorescent substrate. Substrates tested: A. Rhodamine 123, B. Rhodamine 123 + cyclosporin A (1 μM), C. daunomycin, and D. BODIPY-tagged vinblastine. Assays were performed according previously described protocols (Hrycyna et al., 1998), and substrate retention was analyzed by FACS.
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