1 Department of Molecular and Cell Biology, University of Cape Town, South Africa
2 Electron Microscope Unit, University of Cape Town, South Africa
Correspondence
Vernon E. Coyne
vernon{at}science.uct.ac.za
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ABSTRACT |
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The GenBank accession number for the sequence reported in this paper is U61972.
Present address: The Marine Biological Association of the UK, Citadel Hill, Plymouth, UK.
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INTRODUCTION |
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To better understand the reasons for the die-offs of G. gracilis in Saldanha Bay, Jaffray & Coyne (1996) developed an in situ assay to identify putative bacterial pathogens of this macroalga. Of the epiphytic bacteria tested, a positive correlation between an agarolytic phenotype and bacterial pathogenicity was discovered (Jaffray & Coyne, 1996
). Other than the aforementioned example, four other incidences of disease, namely iceice white powdery disease in Eucheuma and Kappaphycus species, the disease in the red alga Rhodella retriculata, the rotten-thallus' syndrome in a Gracilaria species and the white-tip disease of Gracilaria conferta, were all attributed to agarolytic bacteria (Friedlander & Gunkel, 1992
; Largo et al., 1995
; Lavilla-Pitogo, 1992
; Toncheva-Panova & Ivanova, 1997
). However, the role of the agarases in the virulence mechanism of these bacterial pathogens was only hypothesized.
Agar-decomposing bacteria were first isolated by Gran in 1902 (Yaphe, 1957). Consequently, several agarolytic bacterial strains were isolated from marine and other environments. Some of the bacterial isolates have been assigned to the genera Pseudoalteromonas (Akagawa-Matsushita et al., 1992
; Belas et al., 1988
; Groleau & Yaphe, 1977
; Leon et al., 1992
; Potin et al., 1993
; Vera et al., 1998
), Pseudomonas (Ha et al., 1997
; Hofsten & Malmqvist, 1975
; Kong et al., 1997
; Lee et al., 2000
; Malmqvist, 1978
; Nomura et al., 1998
), Cytophaga (Duckworth & Turvey, 1969
; Van der Meulen & Harder, 1975
), Vibrio (Aoki et al., 1990
; Araki et al., 1998
; Fukasawa et al., 1987
; Sugano et al., 1993
) and Streptomyces (Bibb et al., 1987
; Buttner et al., 1987
; Kendall & Cullum, 1984
). Yaphe and co-workers were the first to describe an agar-degrading enzyme system from a marine bacterium (Day & Yaphe, 1975
; Groleau & Yaphe, 1977
). The agar-degrading enzyme system was that of a bacterial isolate classified as Pseudoalteromonas atlantica ATCC 19292T (IAM 12927T) (Gauthier et al., 1995
). The pathway of agar metabolism in this organism involves the initial cleavage of the agarose (alternating 3-O-linked
-D-galactopyranose and 4-O-linked 3,6-anhydro-
-L-galactopyranose) moiety of agar by an endo-acting enzyme,
-Agarase I, yielding neoagaro-oligosaccharides limited by the disaccharide, neoagarobiose unit [O-3,6-anhydro-
-L-galactopyranosyl-(1
3)-o-
-D-galactose], but with predominance of the tetramer, neoagarotetraose [O-3,6-anhydro-
-L-galactopyranosyl-(1
3)-o-
-D-galactopyranosyl-(1
4)-O-3,6-anhydro-
-L-galactopyranosyl-(1
3)-D-galactose], as the major end product. The neoagarotetraose is in turn cleaved at its central
(14) linkages by a neoagarotetraose hydrolase yielding neoagarobiose. However, this enzyme was also shown to be able to degrade species of oligosaccharides larger than neoagarotetraose, and hence it was given the name
-Agarase II. It was shown to hydrolyse agar by an endomechanism to produce neoagaro-oligosaccharides, hexasaccharides, tetrasaccharides and neoagarobiose, with neoagarobiose being the limiting and predominant species. The third and final enzyme in the agar-degrading system of P. atlantica is a neoagarobiose hydrolase that cleaves the
(13) linkage in neoagarobiose to yield the monomeric sugars D-galactose and 3,6-anhydro-L-galactose.
In this study we report the cloning of one of the genes responsible for the agarolytic activity associated with an epiphytic bacterial pathogen of G. gracilis from Saldanha Bay. The enzyme was purified and used as a tool to elucidate its role in the virulence mechanism employed by the bacterium in eliciting disease in G. gracilis.
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METHODS |
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Molecular techniques.
A genomic library was constructed in E. coli HB101, using the plasmid pEcoR251 (Table 1). Genomic DNA was extracted from P. gracilis B9 and partially digested as described by Ausubel et al. (1989)
with Sau3A. The Sau3A DNA restriction fragments were size-fractionated on a 1040 % (w/v) sucrose gradient (Sambrook et al., 1989
). DNA fragments of 10 kb in size were pooled and ligated with T4 ligase into the BglII site of plasmid pEcoR251 at 15 °C overnight (Ausubel et al., 1989
). The recombinant plasmids were recovered by transformation into E. coli HB101 (Dagert & Ehrlich, 1979
). The genomic library was screened for agarolytic activity on LA (Ap) with Gran's Iodine reagent (Groleau & Yaphe, 1977
). Plasmid DNA was isolated using a Nucleobond AX plasmid purification kit (MachereyNagel), according to the manufacturer's instructions, and digested with restriction enzymes obtained from Boehringer Mannheim and Amersham. Agarose gel electrophoresis was performed in Tris/acetate buffer (Ausubel et al., 1989
). Both the Southern hybridization and Heinikoff shortening procedures were followed as described by Ausubel et al. (1989)
. Sequencing was performed with the dideoxynucleotide chain-termination method using either the Sequenase sequencing kit (Amersham Pharmacia) and [
-35S]dATP (Sanger et al., 1977
) or a Thermosequenase cycle-sequencing kit (Amersham) and an ALFexpress automated sequencer (AM version 3.01; Pharmacia Biotech). Sequence data were analysed using DNAMAN version 4.13 (Lynnon BioSoft) and DNASIS software version 2.1 (Hitachi Software Engineering). Homology searches with both DNA and protein sequences were carried out using the BLAST algorithm (Altschul et al., 1990
) provided by the Internet service of the National Centre for Biotechnology Information (http://www.ncbi.nlm.nih.gov/blast/).
Purification of AagA.
Five 1 l E. coli JM109(pDA16) cultures (Table 1) were grown in LB (Ap) for 24 h at 22 °C on an orbital shaker at 100 r.p.m. The cultures were centrifuged and the supernatants collected. The following procedures were performed at 4 °C. The supernatant was adjusted to a final ammonium sulphate saturation of 85 % (w/v) (Englard & Seifter, 1990
). The precipitate was collected by centrifugation, resuspended in 20 mM Tris/HCl buffer (pH 7) and dialysed multiple times against the same buffer. The following procedures were carried out at 20 °C. A column (3x28 cm) of DEAE-Sephadex A-50 (Pharmacia) was activated with 5 M NaCl and then equilibrated with 20 mM Tris/HCl (pH 7) (Rossomando, 1990
). The dialysed concentrate was applied to the column and the column was washed with 20 mM Tris/HCl (pH 7). The proteins were eluted from the column in a stepwise fashion with an increasing NaCl molarity (0·11 M NaCl). Fractions were collected with a Gilson FC 204 Fraction Collector. The resultant pooled total volume was reduced with an Amicon Centricon PM10 filter system. A column (6x1 m) of Sephadex G75 (Pharmacia) that had been equilibrated with 20 mM Tris/HCl (pH 7) was prepared. The active concentrate was applied to the column and fractions were collected. The active fractions were pooled and concentrated. Finally, the active concentrate was dialysed against 10 mM phosphate buffer (pH 7).
Enzymic assays.
Protein concentrations were determined by the Bradford method (Ausubel et al., 1989). Agarase activity was determined by the ferricyanide reducing sugar assay (Park & Johnson, 1949
).
SDS-PAGE and zymograms.
Samples were separated on a 12 % (w/v) SDS-PAGE gel in accordance with the Laemmli method (Ausubel et al., 1989) and detected by silver staining (Sammons et al., 1981
). Zymogram detection of the extracellular agarase was performed as follows: a final concentration of 0·1 % (w/v) agarose was incorporated into the separating gel matrix of a 12 % (w/v) SDS-PAGE gel. After electrophoresis, the gel was soaked in 10 mM phosphate buffer (pH 7) at 22 °C. The buffer was replaced hourly for 3 h before incubation at 37 °C for 12 h. Zones of hydrolysis were visualized by staining the gel with Gran's Iodine.
TLC analysis of agarase hydrolysates.
Purified enzyme (600 ng) was added to 100 µl freshly prepared 1 % (w/v) agarose substrate and 200 µl 20 mM PIPES solution (Yaphe, 1957). Similarly, 600 ng of purified enzyme was added to 50 µl of each of three oligosaccharides: neoagarobiose, neoagarotetraose and neoagarhexaose (Promega) (2·5 µg µl-1 final concentration) and 140 µl 20 mM PIPES solution. The reaction mixes were incubated at 37 °C for 1 h. TLC was performed on Silica gel 60 aluminium foil (Merck) and developed with the solvent n-butanol/acetic acid/water (2 : 1 : 1). The digests were visualized with naphthoresorcinol (Yaphe, 1957
).
Antibody production against purified AagA.
Polyclonal antibodies against purified AagA protein were obtained by immunizing a rabbit with 150 µg AagA, purified from the E. coli JM109(pDA16) transformant, together with Freund's incomplete adjuvant (Ausubel et al., 1989).
Ultrastructure evaluation and colloidal gold immunolabelling.
The procedure described by Dykstra (1993) was adapted for the preparation of pathogenicity assay samples. Thalli (6 mm in length) were washed in a base buffer [PBS, pH 7, 2·4 % (w/v) NaCl] to remove any excess material. The samples were fixed by overnight immersion in 510x sample volume of 2·5 % (w/w) glutaraldehyde in base buffer at 4 °C. The tissues were rinsed twice (5 min each) in base buffer. The samples were post-fixed in 1 % (w/v) osmium tetroxide for 1 h at 22 °C. The tissues were rinsed twice in water (5 min each). Dehydration of the samples was carried out by passing them through the following alcohol dilution series: 30, 50, 70, 80, 90 and 95 % ethanol for 5 min respectively. The samples were then passed through 100 % ethanol for 10 min (twice) and finally through 100 % acetone for 10 min (twice). The samples were then infiltrated with Spurr resin (Spurr, 1969
) as follows. The thalli were placed individually in moulds and covered with Spurr resin. The samples were polymerized in a 60 °C oven for 2 days, after which the polymerized wedges were stored at 22 °C. Ultrathin cross sections of the thallus were obtained with a Leica ultracuts ultramicrotone and mounted on carbon-coated nickel grids using a modification of the method described by Beesley (1989)
. The grids were first floated, section downwards, on PBS containing 1 % BSA (PBS-BSA) for 5 min. The grids were transferred to PBS containing glycine for 3 min and washed twice (1 min each) with PBS-BSA. Duplicate grids were floated on anti-AagA-containing serum obtained from either the first or the fifth bleed for 12 h and washed five times (1 min each) with PBS-BSA containing 0·1 % (w/w) Tween. The grids were washed thrice (1 min each) with PBS-BSA. The grids were then floated on a 1 : 50 dilution of 15 nm gold anti-rabbit probe in PBS-BSA for 2 h. The grids were rinsed five times (1 min each) with PBS-BSA containing 0·1 % (w/w) Tween, followed by three washes (1 min each) with PBS-BSA. The conjugant label complexes were fixed with 1 % (w/w) glutaraldehyde in PBS at 22 °C for 3 min. The grids were rinsed five times (1 min each) in ultrapure water. The grids were stained with 2 % (w/w) uranyl acetate for 10 min and washed five times (1 min each) with ultrapure water. The sections were then stained with a second stain, Reynolds lead citrate, for 5 min and the grids were washed in a stream of ultrapure water for 2 min. The samples were visualized with a JEM-200CX transmission electron microscope (JEOL).
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RESULTS |
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Restriction enzyme mapping of the recombinant plasmid pDA1
Three E. coli HB101 transformants capable of hydrolysing agar were isolated after screening the P. gracilis B9 genomic DNA library for agarase-encoding genes. A preliminary restriction enzyme digest performed on the three agarolytic recombinant plasmids isolated from the three E. coli HB101 transformants revealed that the plasmids were identical (data not shown). A representative was mapped further and the recombinant plasmid was designated pDA1 (Fig. 1a).
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Deletion analysis of pDA1
Since the recombinant plasmid pDA1 harboured a large insert of P. gracilis B9 genomic DNA (pDA1 has an insert of 7·0 kb) it was necessary to identify which region of the P. gracilis B9 DNA fragment was responsible for the agarolytic activity observed in the E. coli HB101 clones. This was achieved by deleting various fragments from pDA1 and visually scoring for agarolytic activity (Fig. 2
). The 1·2 kb HindIIIEcoRI fragment of pDA1 was found to include the P. gracilis agarase gene(s).
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Homology searches
Several databases were searched for homologous sequences to determine the putative identity of the protein encoded by the ORF on the 2·75 kb HindIIIXhoI fragment of pDA012. The ORF was found to have 76 and 85 % identity to the -agarase (dagA, M73783) from Pseudoalteromonas atlantica ATCC 19262T (IAM 12927T) at the DNA and amino acid level, respectively. It also shared a lesser amino acid identity with the two
-agarases (AgaA, AF098954, and AgaB, AF098955) from Cytophaga drobachiensis and the
-agarase (DagA, P07883) from Streptomyces coelicolor A3(2); i.e. 51, 44 and 34 %, respectively. Therefore, it was concluded that the 873 bp ORF encoded a putative
-agarase and was designated aagA.
Purification and characterization of AagA from E. coli JM109(pDA16)
AagA was purified from the extracellular medium of E. coli JM109(pDA16) by using a combination of gel filtration and ion-exchange chromatography (data not shown). The overall yield of AagA was 21·4 % for a purification of 21·7-fold (Table 2). An aliquot of the final concentrate, subjected to SDS-PAGE, exhibited a single band at 30 kDa (Fig. 3
a, lane 1). The zymogram confirmed that the purified protein was an agarolytic enzyme (Fig. 3b
). TLC revealed that AagA hydrolyses
(14) linkages in agarose to predominantly yield neoagarotetraose as the major end product (Fig. 4
, lane 1). In addition, AagA hydrolysed the neoagarohexaose to produce neoagarotetraose and neoagarobiose (Fig. 4
, lane 2). AagA did not hydrolyse neoagarotetraose and neoagarobiose (Fig. 4
, lanes 3 and 4).
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To evaluate the role of the P. gracilis B9 agarase in the infection process, G. gracilis thalli were injected with purified agarase and examined for disease symptoms over a 5 day period. Thalli injected with purified AagA exhibited the largest lesions (10±1 mm). Bleaching in these thalli occurred 2 days post-injection. No bacteria were isolated from AagA-injected thalli after the 5 day incubation period. The injected areas, corresponding to the bleached areas of thalli that had been injected with either P. gracilis B9 or AagA, were washed in a base buffer to remove any excess material and prepared for transmission electron microscopy. A comparison of the cross-sections of the thalli that had been injected with either SSW, P. gracilis B9 or bacterium SS5g revealed no apparent differences in the cell structure. However, a comparison between any of these three cross-sections and that of AagA-injected thalli revealed a clear disruption of the algal cell structure, i.e. the cell walls generally appeared more swollen in comparison to thalli injected with bacterium SS5g (Fig. 5).
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DISCUSSION |
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A P. gracilis B9 genomic library was screened for agarolytic activity by visual inspection of the agar for zones of pitting around E. coli transformants growing on solid media. Agar-digesting E. coli clones were found to harbour a recombinant plasmid designated pDA1. Deletion analysis of pDA1 revealed the location of the gene responsible for agarolytic activity. A BLAST search of the GenBank database showed that the ORF situated in the P. gracilis B9 DNA of pDA012 had sequence identity to a number of -agarases. The
-agarase DagA (M73783) of P. atlantica ATCC 19262T had the greatest similarity to the P. gracilis B9 gene. Therefore, it was concluded that the 873 bp ORF encoded a putative
-agarase and was designated aagA.
A 30 kDa extracellular -agarase was purified from the spent growth medium of the transformant E. coli JM109(pDA16). The size of the purified
-agarase was consistent with the theoretical size predicted for the mature
-agarase. In addition, zymogram analysis confirmed that the purified protein was agarolytic. TLC data confirmed that the purified extracellular agarase was indeed a
(14) agarase; i.e. the agarase hydrolysed the
(14) linkages of agarose to predominately produce neoagarotetraose. However, the
(14) agarase only hydrolysed saccharides larger than neoagarotetraose. This is consistent with the extracellular
-agarase type I enzyme from P. atlantica ATCC 19262T (Day & Yaphe, 1975
; Morrice et al., 1983
).
The polyclonal antibodies raised against the -agarase purified from the E. coli JM109(pDA16) transformant specifically cross-reacted with AagA and the extracellular
-agarase purified from P. gracilis B9. Since a 30 kDa band was the only protein detected in the extracellular extract of P. gracilis B9, it is highly likely that AagA is the only
-agarase secreted into the medium under the growth conditions tested. However, we cannot rule out the possibility that P. gracilis B9 may produce additional extracellular agarases under different growth conditions. For example, when the culture conditions used to grow Vibrio sp. JT0107 were changed, another agarase, designated agarase 0072, was produced by the bacterium (Sugano et al., 1995
).
The relationship observed between disease symptoms exhibited by infected G. gracilis and the agarolytic phenotype of P. gracilis B9 was confirmed. Microscopic examination of cross sections prepared from severely bleached thalli that had been injected with purified AagA revealed disruption of the cell structure. The cell wall appeared to have weakened, i.e. the ultrastructure of the cell wall was lost upon treatment with gluteraldehyde during sample preparation, resulting in a swollen appearance in comparison to cross sections of thalli that had been injected with either SSW, P. gracilis B9 or bacterium SS5g. Immunogold-labelled antibodies localized the -agarase, in situ, to the intercellular matrix of the cell walls of thalli injected with either AagA or P. gracilis B9. In addition, no immunogold-labelled antibodies were detected in the SSW and SS5g sections. There seems to be a direct relationship between the severity of thallus bleaching and the degree of disruption of the fibrillar component of the cell walls. The cellulosic fibrillar cell wall component functions in concert with the mucilaginous agar component of the cell wall to strengthen the thallus (Christiaen et al., 1987
). We hypothesize that
-agarase degradation of the mucilaginous component weakened the overall structure of the cell wall and consequently, resulted in the collapse of the fibrillar component in the AagA-injected thallus. The swollen appearance of the cell wall of AagA-injected thalli could be attributed to an overall loss in cell wall strength of the bleached thalli.
Weinberger et al. (1999) showed that Gracilaria conferta responded with an oxidative burst, a rapid increase in respiration and halogenation when it detected the breakdown products of agar, namely neoagarohexaose and neoagarotetraose. Neoagarohexaose elicited a release of hydrogen peroxide that resulted in an immediate increase in algal brominating activity. Bleached thallus tips appeared a few hours after the addition of neoagarohexaose. These observations are consistent with our results. The end products released as a result of AagA activity on the mucilaginous component of the cell wall could elicit a similar response. Thus, the extensive bleaching that followed injection of pure enzyme (AagA) into the G. gracilis thallus, and the thallus bleaching that occurred as a consequence of secretion of P. gracilis B9 agarase into the thallus of infected G. gracilis, could be due to the macroalga responding to the end products (neoagarohexaose and neoagarotetraose) produced by the extracellular
-agarase of P. gracilis B9 as a consequence of agar degradation.
Even though the central medullary cells were employed as the site of injection in the pathogenicity assay, -agarase was detected in both the medullary and cortical cell walls of thalli that had been injected with either AagA or P. gracilis B9. Since bacteria were only observed at the site of injection, our data suggest that the enzyme spread through the cell wall from the site of injection to other areas of the thallus. Thus, it could be hypothesized that the
-agarase is secreted into the thalli by the bacterial pathogen and becomes associated with the mucilaginous component (agar) of the cell wall. The breakdown products of the hydrolysed agar, namely neoagarohexaose and neoagarotetraose, are then released from the polymer into the surrounding medium, possibly by diffusion stimulated by wave action. Of the agarolytic pathways characterized to date, more than one enzyme has been shown to be required for complete agar hydrolysis, and furthermore, each pathway included both an extra- and intracellular enzyme (Belas et al., 1988
; Potin et al., 1993
). Hence, it is most likely that P. gracilis B9 hydrolyses the oligosaccharides further by means of a cell-bound enzyme(s) that has not yet been identified.
The extracellular -agarase was localized to the G. gracilis cell wall following injection with P. gracilis B9. However, the ultimate effect of the enzyme on the thallus had not yet occurred when the cross sections were prepared for microscopic examination, i.e. complete disruption of the cellulosic fibrillar appearance of the cell wall as a consequence of the degradation of the mucilaginous agar component and severe thallus bleaching. However, when P. gracilis B9-injected thalli were maintained at an incubation temperature of 30 °C, as opposed to 22 °C, the characteristic bleached phenotype was observed (data not shown). The higher incubation temperature results in an increased enzyme activity, which was observed in in vitro enzyme assays (data not shown). Thus, the higher incubation temperature sped up the effect of the enzyme on the thallus. Consequently, we postulate that the bacterial epiphyte P. gracilis B9 becomes pathogenic towards G. gracilis in response to specific environmental conditions.
The immunogold detection technology described in this study could be used for a thorough and detailed investigation of the causes of the disease experienced by G. gracilis at Saldanha Bay. It is difficult to pinpoint the exact factors that cause disease in G. gracilis at Saldanha Bay since the early stages of infection cannot be detected with the naked eye. The symptoms only become visible once the disease has progressed to the point where thallus bleaching and breakage occurs. However, by this time the agar component of the cell wall is degraded and the crop is rendered useless. In situ monitoring of bacterial extracellular -agarases at various times during the G. gracilis growth season, either in natural or raft-cultivated populations at Saldanha Bay, would allow elucidation of environmental cues that result in bacterial infection and the subsequent collapse of G. gracilis populations. Advances in immunocytochemistry have led to the development of non-microscopic techniques such as pregnancy diagnostic kits (Beesley, 1989
). Similarly, an in situ agarase detection kit could be developed to assist Gracilaria farmers in early detection of the onset of disease and thus allow for early harvesting, which would in turn circumvent the complete collapse of the entire Gracilaria crop, thus avoiding financial ruin.
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ACKNOWLEDGEMENTS |
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Received 23 May 2003;
revised 26 June 2003;
accepted 26 June 2003.
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