Detection and identification of ... - Wiley Online Library

The Plant Journal (2007) 50, 95–107

doi: 10.1111/j.1365-313X.2007.03033.x

Detection and identification of rhamnogalacturonan lyase activity in intercellular spaces of expanding cotton cotyledons Radnaa Naran, Margaret L. Pierce and Andrew J. Mort* 246 NRC, Department of Biochemistry and Molecular Biology, Oklahoma State University, Stillwater, OK 74078, USA Received 19 August 2006; revised 14 November 2006; accepted 29 November 2006. *For correspondence (fax +1 405 744 7799; e-mail [email protected]).

Summary Rhamnogalacturonan lyase (RG lyase) activity has been detected and its relative activity measured in vivo during the expansion of cotton (Gossypium hirsutum L.) cotyledons. Rhamnogalacturonan (RG) oligomers labeled with a fluorescent tag were injected into the intercellular spaces of cotton cotyledons and, after incubation, the digested substrate was rinsed out. Enzyme digestion products were detected and identified by capillary zone electrophoresis. Rhamnogalacturonan lyase products were identified as such by co-migration with the digestion products of linear RG oligomers when the oligomers were treated with fungal RG lyase but not when treated with fungal RG hydrolase. In addition, reaction of plant RG lyase digestion products of RG oligomers with I2/KI, which selectively removes unsaturated galactopyranosyluronic acid (GaLap) residues formed at the non-reducing end of the oligomer, converted the plant digestion products into RG oligomers that co-migrated with fungal RG hydrolase products. The activity of the enzyme in the intercellular spaces of cotton cotyledons is very low and could be detected most easily when not >0.03 nmol of substrate was injected in a 0.7-cm2 area and incubated in vivo for 2–6 h. Rhamnogalacturonan lyase activity was the highest in rapidly expanding 3- to 4-day-old cotyledons and gradually decreased during the slow-down in expansion over the next 2–3 days. The RG lyase activity was also detected when the APTS (8-aminopyrene-1,3,6-trisulfonic acid, trisodium salt)-labeled substrates were introduced into intercellular spaces by infiltration instead of injection, indicating that the activity was not induced by wounding or released into the apoplast by cell damage. An exoRG galacturonohydrolase activity was also found, but RG hydrolase and exo-RG rhamnohydrolase were not detected. Keywords: plant cell wall, RG I, RG lyase, RG hydrolase, pectin-degrading enzymes.

Introduction Pectins are one of the quantitatively important constituents of primary plant cell walls of dicots and the non-graminaceous monocots. Four distinct regions of pectic polysaccharides have been extensively studied. They are homogalacturonan (HG), rhamnogalacturonan I (RG I), rhamnogalacturonan II (RG II) and the more recently discovered xylogalacturonan (Schols et al., 1995). Much attention has been paid to plant-produced enzymes that degrade HG and their roles in fruit ripening (Crookes and Grierson, 1983), cell separation and abscission (Meakin and Roberts, 1990; Sander et al., 2001). There has also been some speculation that they may play a role in cell elongation (Domingo et al., 1998) and tissue expansion (Vogel et al., 2002; Zhang, 1998). Many fungi and bacteria produce HGdegrading enzymes, which are thought to be important in ª 2007 The Authors Journal compilation ª 2007 Blackwell Publishing Ltd

plant pathogenesis and in the use of HG as a carbon source in saprophytic growth on plants. With the exception of b-galactosidases acting on sidechains of RG I (Gross et al., 1995), enzymes in plants degrading the regions of pectin other than HG, whether produced by the plant itself or by potential pathogens, have received far less attention. Little is known, too, about changes in RG I during plant growth and development; although, according to a recent model of pectin in which it is the core that holds the other parts of pectin together (Vincken et al., 2003), it may play a central role in determining both the structure and function of cell walls. Rhamnogalacturonan I consists of a repeating disaccharide [galactosyluronic acid rhamnose (GalA - Rha)] backbone with frequent neutral sugar side chains predominantly of linear and branched oligosaccharides composed 95

96 Andrew J. Mort et al. of a-L-arabinofuranosyl (a-L-Araf ) and b-D-galactopyranosyl (b-D)-Galp) residues attached to C-4 of many of the rhamnopyranosyl (Rhap) residues. It has recently been proposed that HG and xylogalacturonans might also be sidechains on the RG I (Vincken et al., 2003). Rhamnogalacturonan-degrading endo-acting enzymes such as RG hydrolase (rhamnogalacturonan a-D-galactopyranosyluronide-(1,2)-a-L-rhamnopyranosyl hydrolase) and RG lyase (rhamnogalacturonan a-L-rhamnopyranosyl-(1,4)a-D-galactopyranosyluronide lyase) have been found in ‘Pectinex’, a crude enzyme preparation from Aspergillus aculeatus, and cloned and expressed in Aspergillus oryzae (Dalboege et al., 1994). The RG lyase activity, first found in A. aculeatus along with the RG hydrolase, had been initially reported as RG hydrolase B (Dalboege et al., 1994). Nelson et al. (1997) suggested that a putative protein, which has an extensive homology with RG hydrolase B, which was later identified to be RG lyase (Mutter et al., 1996), was inexplicably required for normal sexual development of Neurospora crassa. Exo-acting enzymes such as RG rhamnohydrolase and RG galacturonohydrolase, also found in fungi, have been isolated, purified and characterized (Mutter et al., 1994, 1998a). There are two reports of finding putative RG I-degrading activities in plants. Hydrolysis of rhamnogalacturonan by RG hydrolase yields products containing non-reducing terminal rhamnose residues. An assay was developed to detect these residues by hydrolyzing them with a purified rhamnosidase and quantification by GLC of the rhamnose released (Gross et al., 1995). Low amounts of activity were found in tomato, grape and apple fruits. In a survey of the pectolytic enzymes of carrot roots and leaves (Stratilova et al., 1998), RG hydrolase activity was assayed as an increase in reducing groups formed in an RG extracted from flowers of Malva mauritiana L., and RG lyase was assayed as the increase in absorbance at 235 nm, presumed to be from formation of unsaturated GalA (uGalA; D-(4,5)-unsaturated a-D-GalAp). Activity was reported for both enzymes in the roots but only for the hydrolase in the leaves. Neither of these preliminary reports involved characterization of products in enough detail to ensure that what was being assayed was actually the enzyme suspected. When fungal RG lyase was expressed in potato (Solanum tuberosum L.), the transgenic plants produced morphologically altered tubers, and gross changes were observed in the cell wall composition including a decrease in RG I and relative increase in HG (Oomen et al., 2002). The cell wall material was more readily extracted from transgenic plants compared to the wild type. Rhamnogalacturonan lyaseexpressing tubers showed a developmental phenotype, and galactan and arabinan epitopes were greatly reduced and appeared at altered locations in tuber walls. According to the data, wild-type potato tubers have no detectable RG lyase activity.

Lyases cleave via a b-elimination mechanism, generating oligomers with unsaturated uronic acid, e.g. uGalA, at the non-reducing end (Linhardt et al., 1986). Generation of unsaturated uronic acid is usually shown by increased absorption at 235 nm, but the presence of unsaturated products in nanomole quantities is difficult to confirm. In previous work with RG lyase, the presence of uGalA was confirmed by NMR spectroscopy (Mutter et al., 1998b); however, the presence of unsaturated product could not be confirmed in another work (Mutter et al., 1998c). There is a patented method for removing D-(4,5)-unsaturated uronic acid from the non-reducing end of glycosaminoglycans after digestion with heparin lyase (Van Boeckel and Van Dedem, 1990). Using a slight modification of this method, we succeeded in converting oligomers of 1,2-a-L-Rhap and 1,4-a-D-GalAp dimers with uGalA at the non-reducing end and Rha at the reducing end [uG(RG)mR; digestion products of cotyledon RG lyase activity] into oligomers of 1,2-a-LRhap and 1,4-a-D-GalAp dimers with Rha at the reducing end [(RG)mR], like the digestion products of RG hydrolase activity toward oligomers of 1,4-a-D-GalAp and 1,2-a-L-Rhap dimers with Rha at the reducing end (GR oligomers). We report here on the detection and identification of RG lyase activity in cotton cotyledon intercellular spaces. Results Detection and identification of RG lyase activity in cotton cotyledon intercellular spaces When (GR)n oligomers, where n = 7, 8 or 9, pre-labeled with fluorescent 8-aminopyrene-1,3,6-trisulfonic acid, trisodium salt (APTS) tags were injected into the intercellular spaces in cotton cotyledons, degradation of the oligomers was observed by capillary zone electrophoresis (CZE) of intercellular wash fluid (IWF) after 2–6 h of incubation. The product peaks, which were regularly spaced, appeared to belong to a homologous series produced by cleaving the oligomer between repeat disaccharides (Figure 1). To check if the enzyme activity was induced as a result of wounding or was released from inside cells that had been damaged by the injection needle, the fluorescent substrate was vacuum infiltrated in several cases into the intercellular spaces rather than injected, but the same pattern of enzyme digestion was observed. (This method of introduction of the substrate consumed much more of the fluorescent substrate, so was not used routinely.) To ascertain if the activity was from contaminating microorganisms, several plants grown under sterile conditions were tested and found to produce the same degradation of the substrate. We concluded that the enzyme activity was a normal extracellular constituent of healthy cotton cotyledons. Recombinant RG hydrolase expressed in Pichia pastoris, the specificity and action pattern of which have been well

ª 2007 The Authors Journal compilation ª 2007 Blackwell Publishing Ltd, The Plant Journal, (2007), 50, 95–107

Rhamnogalacturonan lyase activity in expanding cotton cotyledons 97

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Figure 1. Electropherograms of APTS-labeled GR-oligomers (a–c) and their partial digests by cotyledon RG lyase (d–f). Peaks were identified based on results presented in subsequent figures. Peaks labeled ‘x’ are (GR)7, present as a mixture in (GR)8, and (RG)6R, derived from (GR)7 by the action of exo-RG galacturonohydrolase.

established (Fu et al., 2001), was used to produce (RG)mR oligomers for comparison with the oligomers produced in the cotyledons. Fungal RG hydrolase needs at least 10 sugar residues, i.e. (GR)5, for activity and cuts between GalA and Rha, producing a number of homologous oligomers with fluorescent label at the reducing end from a labeled (GR)n oligomer: (RG)2R to (RG)n–3R (Fu et al., 2001). In previous work with recombinant RG hydrolase (Fu et al., 2001), amino-naphthalene trisulfonic acid (ANTS) was used to label GR oligomer substrates with a fluorescent tag. Labeling with APTS instead considerably increased the sensitivity of detection, since APTS has more than twice the molar extinction coefficient of ANTS as well as a greater quantum efficiency (Evangelista et al., 1995). Fungal RG hydrolase (RGase) digestion products are designated as the (RG)mR series, and RG lyase digestion products of the same substrate, having uGalA at the non-reducing end and Rha at the reducing end, are designated as uG(RG)mR, m being the number of disaccharide repeating units. For example, from (GR)9 the fungal hydrolase produces five labeled oligomers: (RG)2R up to (RG)6R (Figure 2c). In contrast, in planta digestion yielded some six products [plus (RG)8R arising from exo-RG galacturonohydrolase activity], as shown in Figures 1(f) and 2(b). This pattern of cotyledon enzyme action on linear (GR)n oligomers was observed consistently from (GR)9 and also from (GR)7 (Figure 1d) and (GR)8 oligomers (Figure 1e). In planta enzyme activity has been demonstrated only against RG oligomers of up to 18

residues in length. It seems likely that the enzyme would also act on polymeric RG, but this has not been shown, so it could be an RG oligosaccharide lyase. The products of in vivo digestion and those of digestion by recombinant RG hydrolase of identical GR oligomeric substrates separate from each other on CZE (Figure 2d), indicating that the in planta enzyme activity is different from the fungal RG hydrolase. If the enzyme acting in vivo is RG lyase, it would produce GR oligomers with D-4,5-unsaturated GalA at the nonreducing end. To identify such a product, we looked for a method to convert uG(RG)mR oligomers into (RG)mR – like the products of fungal RGase digestion of GR oligomers – by removing the uGalA. A slight modification of a method that splits off D-(4,5)-unsaturated a-D-GlcAp (uGlcA) from the heparin lyase-digested glycosaminoglycans (Van Boeckel and Van Dedem, 1990) proved to be suitable for this purpose. Figure 3 illustrates the interconversions. Prolonged incubation of in vivo digestion products with I2/KI retained a homologous series of peaks on CZE (Figure 2f). The I2/KI-treated in vivo digestion products separated on CZE from the initial in vivo digestion mixture (Figure 2g), and they had identical retention times on CZE as the products of fungal RGase digestion of the same oligomer (Figure 2h), which strongly suggests that the enzyme acting in intercellular spaces of cotton cotyledons is RG lyase. Confirmation of the enzyme identification as RG lyase came from co-electrophoresis of the products of in vivo


98 Andrew J. Mort et al.

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enzyme digestion of GR oligomers with products using A. aculeatus RG lyase; the product peaks from the fungal RG lyase and cotyledon RG lyase treatments did not separate on CZE (Figure 4d). [I2/KI-treated and untreated fungal RG lyase digestion products separated from each other (Figure 4 g,h) as was seen for the cotyledon digestion products (Figure 2g).] The specificity of the plant RG lyase is a little different from that of the fungal RG lyase purified from A. aculeatus (Mutter et al., 1998c) although they produce the same smallest oligomer. The oligomers produced by fungal RG lyase digestion of unlabeled (GR)10 were identified by matrixassisted laser desorption/ionization time-of-flight mass spectroscopy (MALDI-TOF MS) (see Figures S1 and S2). The smallest oligomer, seen best in the positive linear mode, had an m/z of 667 corresponding to uGRGR+Na+. After

Figure 2. Electropherograms of digests of APTSpre-labeled (GR)9 by cotyledon RG lyase, fungal RGase, and fungal RG lyase, and their co-electrophoresis. (a) APTS-labeled (GR)9. (b, e) Partial digest of (GR)9 by cotyledon RG lyase. (c) Partial digest of (GR)9 by fungal RGase. (d) Mixture of (b) and (c): products of (GR)9 by cotyledon RG lyase separate from fungal RGase products on CZE. (f) Same digest as (e) after I2/KI treatment to remove non-reducing end uGalA from cotyledon RG lyase digestion products. (g) Mixture of (e) and (f): cotyledon RG lyase products separate from their I2/KI-treated products. (h) Mixture of (c) and (f): I2/KI treatment removes uGalA from cotyledon RG lyase products, and they do not separate from fungal RGase products on CZE. (RG)8R in (b) and in (d) to (h) is the product of the action of exo-RG galacturonohydrolase action. Peaks 1 to 6 are uGRGR to uG(RG)6R; 1a to 6a are RGR to (RG)6R; x is an unidentified peak.

labeling of the uGRGR with APTS, it co-migrated with the smallest product from fungal RG lyase digestion of APTS-prelabeled (GR)7. Co-electrophoresis of in planta digestion products with fungal RG lyase products confirmed that uGRGR is also the smallest oligomer produced by cotyledon RG lyase (Figure 4b–d). These results indicate that the enzymes need four sugar residues to the reducing side of the enzyme cut site. The plant enzyme can produce labeled oligomers as long as uG(RG)6R from labeled (GR)9 (Figures 1f, 2b and 4b), whereas the fungal enzyme makes oligomers only up to uG(RG)3R (Figure 4c,e). This shows that the plant enzyme only needs four sugar residues to the non-reducing side of the cut site, but the fungal one requires 10. (See Figure 5 for the digestion patterns of cotyledon RG lyase, A. aculeatus fungal RG lyase and Botrytis cinerea fungal RG hydrolase.)



Figure 3. Fungal RGase (I) and cotyledon RG lyase (II) digestion products of (GR)n oligomer: The digestion products of cotyledon RG lyase can be converted to be the same as fungal RGase products by treatment with I2/KI, which selectively removes non-reducing uGalA.

Are there D-4,5-unsaturated GR oligomers in cotyledon intercellular spaces? In an attempt to find endogenous RG lyase-produced oligosaccharides in the IWF of cotyledons, several aliquots of IWF were labeled with APTS and then analyzed by CZE. At migration times characteristic for oligomers from RG I, there were broad irregular peaks of low intensity. No distinct peak co-migrated with the labeled products we rinsed from cotyledons after incubation with labeled GR oligomers. The lyase activity is not extracted by solutions with a high ionic strength Cotyledon IWF was prepared by infiltration with 50 mM and 200 mM ammonium acetate (NH4Ac), pH 5.2, and with 500 mM NaCl to extract ionically bound RG I-degrading enzymes from the apoplast. The 50-mM extract showed no activity against labeled (GR)8 (Figure 6b). However, 200 mM

NH4Ac, pH 5.2, and 500 mM NaCl (Figure 6c, d) both demonstrated exo-RG galacturonohydrolase activity toward the labeled oligomer. Rhamnogalacturonan lyase activity was not detected in any of the extracts. Since it has been demonstrated by infiltration of RG oligomers that RG lyase activity is present in the apoplast, this finding suggests that the enzyme is more strongly, perhaps covalently, bound to the cell wall. Assay of the RG lyase activity in cotyledons of different ages Estimating the amount, and changes in amount, of enzyme activity in the intercellular spaces required the development of a new assay. Substrate solution, (GR)9, 0.03 nmol in 5 ll of 50 mM NH4Ac buffer, pH 4, was incubated with increasing amounts of fungal RGase for 4 h, and a standard curve of percentage conversion to products vs. enzyme units was prepared. The percentage conversion was calculated by dividing the sum of the modified peak areas for product oligomers by the total of all the modified peak areas (see


100 Andrew J. Mort et al. (GR)9

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Figure 4. Electropherograms of partial digests of (GR)9 by cotyledon and fungal RG lyases and removal of uGalA by I2/KI treatment of the fungal RG lyase digest. (a) APTS-pre-labeled (GR)9. (b) Cotyledon RG lyase digest. (c), (e) Fungal RG lyase digest. (d) Mixture of (b) and (c). (f) Same digest as (e) after I2/KI treatment. (g) 8:4 mixture of (e) and (f). (h) 8:5 mixture of (e) and (f) to show that (RG)mR oligomers migrate more slowly than corresponding uG(RG)mR oligomers except for RGR. (See Experimental procedures for explanation.) Peaks 1 to 6 are uGRGR to uG(RG)6R, 1a to 3a are RGR to (RG)3R.

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Experimental procedures). The assay was linear with respect to the enzyme activity in the range from 20% to 60% conversion of substrate to products. An advantage of using the percentage conversion of substrate to products is that one need not obtain a quantitative recovery from the intercellular spaces so long as the ratio between products and substrate is retained during the preparation of the extract. In an experiment in which a mixture of labeled oligomers was injected, all oligomers were recovered from the intercellular spaces in the same ratio as the initial mixture. The same amount of substrate as was used for the preparation of the standard curve was injected into intercellular spaces of 3- to 7-day-old cotyledons and incubated for 4 h, which gave the percentage conversion of the substrate in the acceptable range. The percentage conversion calculated after in vivo digestion was converted to enzyme units

by comparison with the standard curve. After an initial rise in the very young cotyledons, the amount of RG lyase declined as a smooth function of age (Figure 7). The amount of RG lyase was highest in rapidly expanding 4-day-old cotyledons, and it gradually decreased during the slow down in expansion over the next 2–3 days. To be able to estimate what area of cotyledon was exposed to substrate during the incubation and indicate from what area the enzymic activity was being sampled, 5 ll of solution of APTS-labeled (GR)3, at the same concentration as used for the CZE experiments, was injected into cotyledons. After a 4-h incubation the cotyledons were cut into 3- to 5-mm strips, and the APTS fluorescence was observed by confocal microscopy. As can be seen in Figure 8, the infiltrated area was sufficiently fluorescent to illuminate the outline of cells in the epidermis. There was a sharp drop-off


Rhamnogalacturonan lyase activity in expanding cotton cotyledons 101 (GR)8

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Figure 5. Cleavage patterns of APTS-pre-labeled GR oligomers by cotyledon and fungal RG lyases and by fungal RGase.

in intensity within a few millimeters of the boundary of infiltration – in the first strip outside the boundary, the fluorescence was strong around a vein but not elsewhere. In one strip further removed, the APTS fluorescence was very weak. The results indicate that the injected substrate did not spread far from the infiltrated area. When 5 ll of substrate solution was injected into a cotyledon, the area of infiltration, which varied slightly depending on the age of cotyledon, was 0.7 cm2. Thus, one could calculate the approximate amount of enzyme present in a cotyledon on an area basis as in Figure 7. Discussion We are confident that the results presented here show there is RG lyase activity in the intercellular spaces of expanding cotton cotyledons. Pure fluorescent GR oligomers incubated in the intercellular spaces were converted into shorter oligomers in a time-dependent manner, and most of these oligomers co-migrated on CZE with oligomers generated

X

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Time, min Figure 6. CZE of (GR)8 after incubation with salt-extracted IWF. (a) Control, (GR)8 pre-labeled with APTS. (b) 50 mM NH4Ac-extracted IWF incubated with (GR)8 pre-labeled with APTS. (c) 200 mM NH4Ac-extracted IWF incubated with (GR)8 pre-labeled with APTS. (d) 500 mM NaCl-extracted IWF incubated with (GR)8 pre-labeled with APTS. All incubations were carried out under same conditions with equal quantities of the substrate and IWF. In (c) and (d), (RG)7R is the product of exo-RG galacturonohydrolase digestion of (GR)8. As seen from the CZE, 200 mM NH4Ac and 500 mM NaCl were able to extract ionically bound exo-RG galacturonohydrolase from the apoplast. Broadening and shifting of peaks, especially in (c) and (d), is due to a high salt concentration in the analysis mixtures. Peaks labeled ‘x’ are unidentified.

from the same GR oligomer digested with an authentic fungal RG lyase. Both the in vivo and in vitro generated oligomers could be converted by I2/KI treatment into


102 Andrew J. Mort et al. 5.5 10 –7

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Figure 7. Amounts of RG lyase detected in expanding cotton cotyledons. Error bars represent standard deviation of three in planta digestion experiments for each day post-emergence.

Figure 8. Confocal laser scanning microscope images of a cotyledon infiltrated with (GR)3 labeled with a fluorescent tag: images, 4 h after injection of the oligomer, from the original area of infiltration (a), and at approximately 2 mm (b) or 6 mm (c), from the boundary of infiltration.

oligosaccharides that co-migrate with oligosaccharides generated by digestion of the starting fluorescent oligosaccharide by an authentic RG hydrolase. The approach of using I2/KI treatment to cause a change in the electrophoretic mobility of only those oligosaccharides containing an unsaturated uronic acid residue should be applicable to the detection of pectate and pectin lyases. There are only a few reports of finding pectate lyase activity in plants (Domingo et al., 1998; Marin-Rodriguez et al., 2003; Payasi and Sanwal, 2003; Stratilova et al., 1998), although it has been predicted that Arabidopsis codes for 28 pectate and pectin lyases (Henrissat et al., 2001). Use of an APTS-labeled galacturonic acid oligomer with CZE for analysis and the I2/KI method for confirmation should allow detection of pectate lyases in very small amounts of tissue. For example, it should be possible to confirm whether Arabidopsis pollen, in which two messages resembling those of bacterial pectate lyases have been found (Wing et al., 1989), produces pectate lyase. It should also be possible to use fluorescent galacturonic acid oligomers, injected as a substrate in leaves, to determine whether the powdery mildew-resistant Arabidopsis mutant pmr6 has lower extracellular pectate lyase levels than wild type as predicted by Vogel et al. (2002). In an approach complementary to this, Bauer et al. (2006) have used cloned fungal enzymes of known activities on extracted cell wall polymers coupled with CZE of APTS-labeled oligosaccharide products to learn that the irx9 mutant of Arabidopsis has reduced xylan content. Those results, plus others, led the authors to infer that IRX9 may be a xylan synthase. Identification of RG lyase activity in cotton cotyledons proved easier than detecting RG lyase products from an endogenous substrate. No distinct RG oligomers were found in IWF. The objective of identifying a peak representing RG lyase products in IWF is made difficult because RG I products are not the only oligomers that might be there. There are very few reports on the detection and identification of native oligosaccharides derived from structural polysaccharides in plant intercellular spaces, but all of these involve finding fragments of the much simpler HG. Galacturonic acid (GalA)-containing oligomers were detected in extracts of tomato fruit during extensive autolysis during ripening (Melotto et al., 1994), and after inoculation with the pathogen B. cinerea (An et al., 2005). Zhang (1998) found high levels (tens of milli-units cm)2 of cotyledon) of endopolygalacturonase in the intercellular spaces of cotton cotyledons; however, after extensive fractionation of IWF from a large number of cotton cotyledons, Miranda (1993) detected a series of GalA oligomers at only picomol cm)2 levels. By comparison, RG lyase was found at 0.2–0.5 microunit cm)2, 1/10 000th the level of endo-polygalacturonase. Until very recently there were no reports of RG lyase genes or activity in any plant other than potatoes in which a fungal lyase gene had been introduced (Oomen et al., 2002). However, we found seven sequences in the Arabid-


Rhamnogalacturonan lyase activity in expanding cotton cotyledons 103 opsis genome that gave a TBLASTN hit with a low bit score of 32–37 to the fungal lyase sequence from A. aculeatus. The CAZY website (http://afmb.cnrs-mrs.fr/CAZY/) predicts that the same seven Arabidopsis sequences are family 4 RG lyases (Coutinho and Henrissat, 1999). The more recently characterized RG lyase from Erwinia chrysanthemi shows much higher homology to the Arabidopsis putative RG lyases, with bit scores of between 108 and 156 for the various members. For all but one of the genes there is evidence that they are expressed, being represented in EST libraries or by cDNA entries in GenBank. Three of the seven genes appear to contain a secretion signal sequence. Curiously, four do not, suggesting that some of the sequences code for lyases whose substrates (probably not RG) are cytoplasmic or that the signal peptide prediction programs are not perfect. A search of dbEST reveals that very similar genes are expressed throughout the angiosperms, from poplar to barley, to tomato. Thus, it is quite likely that RG lyases are important for the wellbeing of all plants. No genes for RG hydrolases have been predicted in Arabidopsis, and we did not find any such activity in cotton cotyledons. The finding of exo-RG galacturonohydrolase activity in cotton intercellular spaces raises some interesting questions. If the plant uses a lyase to cleave the RG what would be the source of substrate for the exo-enzyme? Does the exo-enzyme also work on uGalA? After the removal of a GalA from an RG oligomer, how would the rhamnose be removed since there do not appear to be homologs of fungal or bacterial rhamnosidases in Arabidopsis (Henrissat et al., 2001)? Unfortunately, there are no sequence data on the known fungal exo-RG galacturonohydrolase, so prediction of a homolog in the Arabidopsis genome is not yet possible. The enzyme unsaturated rhamnogalacturonyl hydrolase, which hydrolyzes the unsaturated GalA from RG lyase products, has recently been identified in Bacillus subtilis (Itoh et al., 2006). Such an enzyme has not been reported in plants, and a TBLASTN search of plant sequences yielded a good match only in rice with a bit score of 169. Although the levels of RG lyase activity in cotton cotyledons are much lower than those of endo-polygalacturonase (Figure 7; Zhang, 1998), as mentioned above, both enzymes show their maximum activity in the very young cotyledons, 3 or 4 days after emergence from the soil. This is the time of most rapid expansion (Zhang, 1998), and the rates of both enzyme activities and of expansion all drop in parallel beyond 4 days. The smooth curve of RG lyase activity as a function of cotyledon age, the fairly similar curve of endo-polygalacturonase activity determined from independent experiments, and the correlation of these activities with rate of leaf expansion, all argue against wound activation as a cause of the enzyme activity observed. What may be the role of RG lyase during leaf growth? If RG really is the backbone to which

other regions of pectins attach, then an enzyme cleaving it could cause dramatic effects on the cohesion of the pectin network. This may be necessary during cell wall expansion. Experimental procedures Materials Citrus Pectin was purchased from Sigma (http://www. sigmaaldrich.com/), APTS was from Molecular Probes (http:// probes.invitrogen.com), HPLC grade ammonium acetate and acetonitrile were from Fischer Scientific (http://www.fishersci.com/). All solvents used were filtered through a 0.45-lm nylon filter and degassed in a sonicating bath under vacuum prior to use. Commercial polysaccharide-degrading enzymes were from Megazyme (http://secure.megazyme.com/). The fungal RG hydrolase from B. cinerea was cloned and expressed in P. pastoris (Fu et al., 2001). The fungal RG lyase from A. aculeatus (Mutter et al., 1998c) was kindly provided by Dr Gerrit Beldman from Wageningen Agricultural University, Netherlands. The fungal RG lyase from Aspergillus nidulans (AN6395.2) was cloned and expressed in P. pastoris (Bauer et al., 2006). For ultrafiltration procedures, a device from Amicon and filters YM10 or YM30 (Millipore Corp., http://www.millipore.com/) were used. Anion-exchange and size-exclusion HPLC were performed on Dionex PA1 (Dionex, http://www1.dionex.com/) and Toyopearl HW-40s (Supelco; http://www.sigmaaldrich.com/Brands/supelco_ Home.html) columns, respectively.

(GR)n substrate preparation Linear rhamnogalacturonan oligomers, (GR)n, used for assay were 12–20 residues long with GalA at the non-reducing end and Rha at the reducing end and were produced from Citrus Pectin (Sigma), which contains approximately 10% RG (Zhan et al., 1998). The pectin was saponified, digested with endo-polygalacturonase (EPG) and ultrafiltered (YM30 membrane) to obtain a high-molecularweight fraction of predominantly RG. Rhamnogalacturonan oligomers were produced by partial hydrolysis of the highmolecular-weight fraction with 2 M trifluoroacetic acid at 80C for 4 h and dialysis through 1000 molecular weight (MW) cut-off membrane, followed by separation of the retentate on a Dionex PA1 anion-exchange column. The partial acid hydrolysis preferentially cleaves Rha–GalA glycosidic linkages of the repeating Rha–GalA disaccharide backbone of the polysaccharide and cuts off the Araand Gal-containing side chains. Thus, the resulting oligomers have GalA at their non-reducing ends and Rha at the reducing ends. They are designated as (GR)n, where n is the number of disaccharide repeats. Individual oligomers, generated by separation on a PA1 column, were identified by NMR and MALDI-TOF MS. As determined by MALDI-TOF MS, (GR)7 has MW of 2272.84, (GR)8, 2595.45 and (GR)9, 2917.10 in negative linear mode.

MALDI-TOF MS Oligosaccharides [0.1–0.2 nmol or 300–600 ng (GR)9] were dissolved in 5–10 ll of nanopure water. From the sample solution 0.5 ll was loaded on top of the matrix – equal volumes of 10% solution of 2¢,4¢,6¢-trihydroxyacetophenone (THAP) in methanol, 1.5% solution of nitrocellulose in acetone/2-propanol, 1:1 mixture,


104 Andrew J. Mort et al. and a 20 mM solution of ammonium citrate – and air-dried on the sample plate. Prior to analysis, sample and matrix solutions were desalted with Dowex 50W 50 · 8–200 (NH4+ form; Korner et al., 1998). Spectra were obtained on a Perseptive Biosystems (http:// www.appliedbiosystems.com/) MALDI-TOF Voyager De-Pro mass spectrometer in the negative ion reflector or linear mode.

NMR spectroscopy 1 H NMR, total correlation spectroscopy (TOCSY) and heteronuclear multiple-quantum correlation (HMQC) spectra of samples in D2O were recorded at 25C on a Varian Inova 600 NMR spectrometer (http://www.varianinc.com) using the standard pulse sequences with water pre-saturation.

Amino-pyrene trisulfonic acid (APTS) derivatization of (GR)n The purified individual oligomers (GR)5–(GR)10 were derivatized with APTS by reductive amination as described in Evangelista et al. (1995). Salts and excess labeling reagent were removed from the labeled oligosaccharide by passing through a Toyopearl HW-40S (http://www.toyopearl.com) (100 · 10 mm) gel filtration column eluted with the mixture of 25% acetonitrile and 75% 50 mM ammonium acetate buffer, pH 5.2. To determine the concentration of fluorescent oligosaccharides, we measured A455 nm of solutions of labeled substrates, and concentrations were calculated using molar absorptivity for APTS–sugar adducts as given in Evangelista et al. (1995). Purified substrates were dissolved in nanopure water to make final a concentration suitable for in vivo injection.

Capillary zone electrophoresis (CZE) Capillary zone electrophoresis was performed on a BioFocus 2000 (Bio-Rad Laboratories, http://www.bio-rad.com/) with laser-induced fluorescence detection. A fused-silica capillary (TSP050375, Polymicro Technologies, http://www.polymicro.com/) of internal diameter 50 lm and length 31 cm was used as the separation column for oligosaccharides. The samples were injected by application of 4.5 lb in)2 of helium pressure for 0.22 sec. Electrophoresis conditions were 15 kV/70–100 lA with the cathode at the inlet, 0.1 M sodium phosphate, pH 2.5, as running buffer, and a controlled temperature of 20C. The capillary was rinsed with 1 M NaOH followed by running buffer with a dip-cycle to prevent carryover after injection. Oligomers labeled with APTS were excited at 488 nm and emission was collected through a 520-nm band pass filter. The migration rates of the various oligomers were influenced by the ionic strength of the sample and the age of the capillary. Thus, for positive identification of an oligomer, it was necessary to show co-migration with a known oligomer admixed to the sample. Co-electrophoresis of products from in planta digestion of (GR)9 with its fungal RGase digestion products or in planta digestion products with their I2/KI-treated derivatives revealed that (RG)mR oligomers migrate more slowly than uG(RG)mR oligomers except for RGR, which moves faster than uGRGR (see Figures 2d,g and 4 g,h). At pH 2.5, neutral oligosaccharides labeled with APTS have all their charge from the three sulfonic acid groups of the label. At this pH the silicic acid groups on the surface of the capillary are protonated, so there is no electro-osmotic flow. Thus the migration rates of the labeled oligosaccharides are inversely proportional to their hydrodynamic radius. However, at pH 2.5, galacturonic acid residues are partially negatively charged causing galacturonic acid-containing oligomers to have a higher migration rate than neutral oligosaccha-

rides of equivalent size. Unsaturated uronic acids have an even lower pKa than galacturonic acid, 3.10 compared with 3.51 (Kohn and Kovac, 1978), leading to a greater charge on the lyase products.

Plants for in vivo injection and infiltration Plants of the Acala 44E line of upland cotton (Gossypium hirsutum L.) were grown in flats or clay pots in Jiffy Mix Plus in a Conviron E-15 growth chamber (http://www.conviron.com/) under conditions described elsewhere (Pierce et al., 1993). The same line was planted and grown under sterile conditions on agar to ensure the enzyme activity detected was not from bacteria or fungi.

Enzyme activity in vivo A pot with cotton plants 3–6 days post-emergence was taken from the growth chamber in the middle of the photoperiod. About 5 ll (0.03 nmol) of APTS-labeled oligomer was injected into the intercellular space of a cotyledon by using a gas-tight syringe (10 ll, 1701 RNFS; Alltech Associates, Inc.; http://www.alltechweb.com/) with a fused silica needle of 0.19 mm outside diameter (J&W Scientific Inc.; http://www.cobertassoc.com/j&w_agilent_gc_cap_ col.htm). The plants were put back in the growth chamber after injection. The cotyledons were excised at 2, 3, 4, 5 and 6 h after injection, and intercellular wash fluid (IWF) was obtained as described in Zhang et al. (1996). Cotyledon intercellular space enzymes, which might be present in the IWF, were inactivated by heating at 80C for 10 min. After centrifugation at 16 000 g for 3 min, the IWF was analyzed by CZE. Fluorescent substrates and buffers were infiltrated by applying a water aspirator to the Erlenmeyer flask in which cotyledons were placed in a buffer or a substrate solution. Injection of substrates was preferred over infiltration since the latter method requires an amount of substrate that is practically difficult to obtain.

Enzymic digestions in vitro One microliter of 15, 20, 25, 30, 35 ng ll)1 solutions of freeze-dried fungal RGase medium, the activity of which was 0.015 U mg)1, was added to 5 ll solution of 0.03 nmol APTS-labeled (GR)9 substrate in 50 mM ammonium acetate buffer, pH 4. Three replicates were used for each amount of enzyme. After a 4-h incubation at 30C, the enzyme was heat-inactivated, and the distribution of fluorescence between products and unreacted substrate was determined by CZE with laser-induced fluorescence detection. The extent of the digestion, whether in vivo or in vitro, was estimated as the percentage of the initial substrate that had been converted to product by dividing the sum of ‘modified peak areas’ of the product peaks (relevant to a particular enzyme) by the sum of all ‘modified peak areas’. (Distinguishing unreacted substrate from the product of exo-RG galacturonohydrolase was therefore not necessary for calculating the RG lyase activity.) The modified peak areas were defined as peak areas divided by their migration times to take into account the increased length of time spent in the detection window with decreasing migration rate. A standard curve was prepared.

I2/KI removal of non-reducing end uGalA residue from fungal RG lyase digestion products Approximately 400 lg of unlabeled (GR)10 was digested with cloned A. nidulans RG lyase and dissolved in 200 ll of D2O. After obtaining



(a)

(b)

(c)

hydrolase from RG (Colquhoun et al., 1990), for GRGR produced by partial acid hydrolysis of RG (Zhan et al., 1998) and for uG(RG)2R produced from RG using fungal RG lyase (Mutter et al., 1998b). After digestion of the oligosaccharide with the fungal RG lyase, the spectrum of the sample (Figure 9b) showed several new signals indicative of breakdown of the oligosaccharide via an elimination reaction. As identified by Mutter et al., (1998b) the peak at 5.8 p.p.m. is indicative of the H-4 of a 4,5-unsaturated GalA, and the peak at 5.32 p.p.m. is from H-1 of the Rha to which the uGalA is attached. The peak at 5.14 p.p.m. is from H-1 of the uGalA, and the peak at 4.31 p.p.m. is from H-2 of the Rha to which the uGalA is linked. After the I2/KI treatment none of the peaks indicative of the presence of the uGalA at the non-reducing end of the oligosaccharides were present (Figure 9c). Most of the signals at chemical shifts indicative of a non-reducing terminal Rha overlap with those from Rha in other linkages. However, signal intensity was considerably increased where one would expect H-2 of a non-reducing end Rha, and a signal completely attributable to H-6 of non-reducing terminal Rha appeared at 1.25 p.p.m. In the TOCSY spectrum (data not shown) the peak at 5.22 p.p.m. showed clear correlations to cross-peaks at both 4.05 p.p.m., where H-2 of non-reducing Rha, is expected and 3.97 p.p.m., where H-2 of reducing a-Rha should be. In the HMQC spectrum (data not shown) there were two carbon correlations to the proton chemical shift of 5.22 p.p.m. – one at 101.44 p.p.m., the same as reported by Colquhoun et al. (1990) for H-1 C-1 of nonreducing Rha, and the other at 92.39 p.p.m. consistent with it being from the reducing Rha H-1 C-1. At the proton chemical shift of 4.05 p.p.m. there were two carbon-correlated signals. The one at 71.25 p.p.m. was at the position reported by Colquhoun et al. (1990) for C-2 of non-reducing terminal Rha and the other was at 80.76 p.p.m., consistent with it corresponding to C-2 of a Rha residue with a sugar linked to it through C-2. From these results we can deduce that the uGalA residue produced by the action of the lyase was removed by the I2/KI treatment leaving a non-reducing terminal Rha at the end of the oligosaccharides.

I2/KI treatment of in vivo digestion products

Figure 9. 1H NMR spectra of (GR)10 and its fungal RG lyase digest before and after I2/KI treatment (all in D2O). (a) 1H NMR spectrum of (GR)10. (b) Spectrum of RG lyase digest of (GR)10. (c) Spectrum of RG lyase digest of (GR)10 after I2/KI treatment. NR, non-reducing end; R, reducing end. The region between 1.2 and 1.35 p.p.m. is expanded in the insets for visualization of the indicated peaks.

the 1H NMR spectrum, the digestion mixture was treated with 200 ll of 0.036 M I2 in 2% solution of KI/25 mM ammonium acetate, pH 4.5 for 48 h in the dark. The mixture was freeze dried, after which the dry residue was redissolved in D2O for NMR spectroscopy. The 1D NMR spectrum of the sample at each step was obtained, and both TOCSY and HMQC 2D spectra of the I2/KI-treated sample were collected. Other than lyophylization and resuspension in D2O, no additional sample treatment was done. Thus, after the digestion and the chemical treatment, all non-volatile products were together in the NMR tube. The peaks observed in the 1H NMR spectrum of the (GR)10 oligomer (Figure 9a) can be assigned quite readily from a combination of literature values for RGRG produced by RG

0.024 M solution of I2 in 2% solution of KI in 25 mM ammonium acetate, pH 4.5 (10 ll), was added to IWF (20 ll) containing products of in vivo digestion of a GRn oligomer. The mixture was incubated at room temperature in the dark for 48 h until the reaction was complete. The mixture was then directly analyzed by CZE.

Cotyledon IWF preparation Extracts of ionically bound enzymes were prepared from cotyledons by infiltration of cotyledon intercellular space with 50 mM or 200 mM NH4Ac, pH 5.2, or 500 mM NaCl and recovering IWF by centrifugation of cotyledons at 1400 g.

Confocal laser scanning microscopy Confocal laser scanning microscopy was carried out on a Leica SP2 confocal laser scanning microscope (http://www.leicamicrosystems.com/) using excitation with an argon laser at 488 nm. The emission wavelength was tuned to 520 nm.

Acknowledgements This work was supported by USDA grant USDA/NRICGP 98 01780 and DOE grant DE-FG02-96ER20215 and has been approved for


106 Andrew J. Mort et al. publication by the Director of the Oklahoma Agricultural Experiment Station. We thank Ms Phoebe Doss at the OSU CLSM center.

Supplementary material The following supplementary material is available for this article online: Figure S1. Positive ion, linear mode MALDI TOF MS spectrum of GR10 digested with cloned RG lyase from A. nidulans. Peaks were assigned as follows: 667.21 (uGRGR+Na)+, 683.24 (uGRGR+K)+, 989.16 (uGRGRGR+Na)+, 1005.15 (uGRGRGR+K)+, 1007.07 1311.15 (uGRGRGRGR+Na)+, 1329.13 (GRGRGR+Na)+, + (GRGRGRGR+Na) , 1633.13 (uGRGRGRGRGR+Na)+ and 1651.97 (GRGRGRGRGR+Na)+. The series of peaks spaced 44 mass units apart from 1013 upward are probably from a mixture of polyethylene glycols. The peaks at 1795.08 and 1813.02 are probably from (uGRGRGRGRGR+Na)+ with a single galactose sidechain and the saturated form. The later pair of peaks (the first at 2117.38) probably represents the respective galactosylated oligomers one RG repeat longer. Figure S2. Negative ion, reflector mode MALDI TOF MS spectrum of GR10 digested with cloned RG lyase from A. nidulans. Peaks were assigned as follows: 643.02 (uGRGR-H)), 965.10 (uGRGRGR-H)), 1287.24 (uGRGRGRGR-H)), 1305.22 (GRGRGRGR-H)), 1609.21 (uGRGRGRGRGR-H)) and 1627.17 (GRGRGRGRGR-H)). This material is available as part of the online article from http:// www.blackwell-synergy.com.

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