Chitinolytic activity of Pseudomonas fluorescens isolates from barley and sugar beet rhizosphere

Abstract

Twelve isolates of Pseudomonas fluorescens, isolated from barley and sugar beet rhizosphere as antagonists towards the plant pathogenic microfungi Rhizoctonia solani and Pythium ultimum, showed chitinolytic activity in batch cultures when grown in media without exogenous chitin. Enzyme tests in cultures demonstrated a complete array of chitinolytic enzymes. Endochitinase and chitobiosidase activities seemed to be tightly coupled in the isolates, while there was no relationship to N-acetyl-glucosaminidase activity. Endochitinase activity, showing hydrolysis of chitin polymers, was found to be extracellular in all isolates, although the most active isolates also retained a cell-bound fraction. Isoelectric focusing gel electrophoresis of supernatants containing extracellular enzyme activity showed that all isolates produced one native endochitinase in logarithmic phase. This enzyme was subsequently modified into several isozymes by extracellular processing as the cultures aged in stationary phase. The 12 isolates could be grouped into seven isozymic patterns. Detailed studies of three selected isolates showed that extracellular release of endochitinase activity also took place in stationary phase. The results, however, indicated that in stationary phase regulation of both the overall production of native enzyme and the subsequent formation of isozymes were different among the P. fluorescens isolates.

Chitinolytic activity, Endochitinase, Isozyme pattern, Pseudomonas fluorescens

1 Introduction

Chitin consists of unbranched chains of β-1,4-linked N-acetyl-D-glucosamine (GlcNAc) residues and is part of the cell wall structure in invertebrates, protozoa, fungi and certain groups of algae [1,2]. Degradation of chitin is an important feature in the global recycling of carbon and nitrogen and is primarily a microbial process [2]. Chitinolytic activity may consist of several enzymes hydrolysing β-1,4-bonds between N-acetyl-glucosamine residues: (1) Chitinase (1,4-β-poly-N-acetyl-glucosaminidase; EC 3.2.1.14) or endochitinase (which is the term used in this study), hydrolysing randomly along the chitin polymer and releasing oligomers. (2) Exochitinase (exo-N,N′-diacetylchitobiohydrolase), or chitobiosidase (term used in this study), hydrolysing at the terminal ends of the chitin polymer or shorter oligomers and releasing chitobiose [3]. (3) N-acetyl-glucosaminidase (EC 3.2.1.30), or NAGase (term used in this study), hydrolysing short oligomers, typically chitobiose dimer units and releasing N-acetyl-glucosamine.

Chitinolytic activity is widespread among soil bacteria [2,4]. Chitinolytic activity has been studied intensively in Gram-positive Bacillus circulans[5] and Bacillus licheniformis[6], each harbouring five different isozymes of chitinolytic activity. Among Gram-negative bacteria, Serratia marcescens[7], Enterobacter agglomerans[8], Vibrio harveyi[9] and Alteromonas sp. [10] all contain at least four different isozymes of chitinolytic activity. The present study reports on chitinolytic activity in rhizosphere isolates of Pseudomonas fluorescens. Among the pseudomonads, only Pseudomonas aeruginosa has been studied in detail and found to contain two different isozymes of endochitinase activity [11]. Pseudomonas stutzeri[12,13] and P. fluorescens[14] have been found to produce chitinolytic activity in batch growth cultures, but the enzyme activity has not been characterised in detail.

Chitinolytic activity produced by microfungi and bacteria is a potential mechanism for biological control of plant pathogenic fungi with chitinous cell walls, e.g. Rhizoctonia solani. The antagonistic activity of microbial endochitinases against the root pathogen R. solani is thus a well-documented example of biological control in greenhouse or field experiments [7,8,15,16]. Among the pseudomonads little effort has yet been made to document the role of chitinolytic activity in biological control, although one study of P. stutzeri suggested that this activity was involved in fungal growth inhibition [13].

In a recent study, we reported that several P. fluorescens isolates from sugar beet rhizosphere showed significant antagonism against R. solani by their production of endochitinase activity or antibiotics [17]. However, production of endochitinase activity by the P. fluorescens isolates was variable when assayed by clearing zones on agar plates of different medium composition [17]. The objective of the present study was to further characterise the production of chitinolytic activity in batch cultures of P. fluorescens isolates selected as potential biocontrol agents in rhizosphere. The ecological perspectives of the observations on chitinolytic activities in P. fluorescens isolates are discussed.

2 Materials and methods

2.1 P. fluorescens isolates

A collection of P. fluorescens isolates from barley and sugar beet rhizosphere was selected by their in vitro inhibition of the fungal plant pathogens Pythium ultimum and R. solani as described previously [17]. The isolates were tentatively identified by a combination of both API NE20 (bioMérieux, Marcy-l'Etoile, France) and biochemical tests [17] and characterised as chitinase-positive using a chromogenic spot test [17]. Two chitinase-positive reference strains, P. fluorescens BL915 [14] and P. fluorescens CHA0 [18], reported to control fungal pathogens, were included. Table 1 lists all isolates including their affiliation to biovar based on biochemical characteristics. All isolates were maintained for storage in (50% v/v) glycerol at −80°C.

1

Pseudomonas sp. isolates used in this study

Strain	Source	Species and biovar	Reference
BL915	Soil	P. fluorescens bv. I	[14]
DR54	Sugar beet rhizosphere	P. fluorescens bv. I	[17]
MN222	Barley rhizosphere	P. fluorescens bv. I	This study
DR2	Sugar beet rhizosphere	P. fluorescens bv. III	[17]
DR3	Sugar beet rhizosphere	P. fluorescens bv. III	[17]
PS1	Sugar beet rhizosphere	P. fluorescens bv. III	[17]
PS21	Sugar beet rhizosphere	P. fluorescens bv. III	[17]
CHA0	Tobacco	P. fluorescens bv. V	[18]
PS7	Sugar beet rhizosphere	P. fluorescens bv. VI	[17]
96.578	Sugar beet rhizosphere	P. fluorescens bv. VI	This study
MN272	Barley rhizosphere	P. fluorescens bv. VI	This study
MN370	Barley rhizosphere	P. fluorescens bv. VI	This study

2.2 Batch cultures and incubations

In an initial survey for occurrence of chitinolytic activity in the P. fluorescens isolates, cells were grown for 3 days at 28°C in 25 ml Tryptic Soy Broth (TSB; Difco) on a rotary shaker (200 rpm). Growth was determined by optical density measurements (OD, 600 nm) and chitinolytic activity (see below) was determined in cellular and supernatant fractions after a centrifugation at 7000×g for 7 min at 20°C. The cellular fraction was washed in 0.9% NaCl and resuspended in 10% TSB medium without further concentration. Samples were kept at −20°C for later analysis of enzyme activity and for isoelectric focusing gel electrophoresis (see below). Results were obtained as the mean of three independent replicates.

Endochitinase production was studied in three isolates in detail in TSB medium (30 ml) during 3 days of incubation, representing a full growth cycle of the isolates in batch cultures. The cultures were inoculated with cells, which had been precultured in 25 ml TSB and washed once in 0.9% NaCl, to an OD_600nm of 0.1. The cultures were incubated at 28°C on a rotary shaker (200 rpm). A number of samples (2 ml) were taken at intervals and growth was followed by OD measurements. The supernatant samples were stored at −20°C for subsequent analysis of endochitinase activity and formation of isozymes. Results were obtained as the mean of three independent replicates. To test the stability of endochitinase activity, a control experiment was performed with filter-sterilised (micro-centrifuge tube filter, 0.2 μm, Whatman International, UK) supernatants, taken from an early growth stage (t=15 h) of a batch culture and further incubated at 28°C for up to 3 days. Samples were retrieved at different time intervals to test for endochitinase activity and formation of isozymes.

2.3 Assays of chitinolytic activity

Four different assays using chromogenic or fluorogenic (4-methylumbelloferone; MUF) substrate analogues were used for detailed investigation of the production of chitinolytic enzymes. Endochitinase activity was determined by the use of Carboxy Methyl-chitin-Remazol Brilliant Violet (CM-chitin-RBV; Loewe Biochemica GmbH, Germany) and the (GlcNAc)₄ analogue, MUF-chitotrioside, as substrate. Chitobiosidase activity was determined using the (GlcNAc)₃ analogue, MUF-chitobioside, as substrate. Finally, NAGase activity was determined using the (GlcNAc)₂ analogue, MUF-N-acetyl-glucosaminide, as substrate (all MUF substrates from Sigma).

The chromogenic CM-chitin-RBV substrate was used for a quantitative assay of endochitinase activity in cellular and supernatant fractions of the batch cultures. For this assay a reaction temperature of 50°C was chosen, based on the temperature optimum of approximately 50°C and the high thermostability observed in three representative isolates (PS7, 96.578, DR54) (data not shown). This relatively high temperature optimum is in accordance with observations made for other bacterial endochitinases, e.g. 40–50°C in P. aeruginosa[11], and 50–60°C in S. marcescens[19]. In the assay, a reaction pH of 5.0 was chosen because initial experiments with the isolates indicated a constant optimum at pH 5 and an occasional optimum at pH 8 (data not shown). This slightly acidic pH optimum further compares well to most bacterial chitinases [19,20]. Endochitinase activity with both acidic and alkaline pH optima has been reported [6], and pH optimum in P. aeruginosa was reported to be pH 7–8 [11]. The standard endochitinase assay mixture consisted of a sample (100 μl), Na-acetate buffer (200 μl; 0,05 M; pH 5.0) and CM-chitin-RBV solution (100 μl; 2 mg ml⁻¹) in a water bath at 50°C for 2 min. Buffer and substrate solutions were mixed and preheated for 5 min before the sample was added. The reaction was terminated by the addition of HCl (100 μl; 2 M), causing precipitation of non-degraded substrate. The assay mixture was then cooled on ice for approximately 10 min and centrifuged at 15 000×g at 4°C for 5 min. A sample (350 μl) of the supernatant was finally transferred to wells of a microtitre plate (96 wells, Nunc, Denmark) and measured photometrically at 540 nm in a microplate reader (Microplate Bio-kinetics Reader, Bio-tek Instruments, USA). A culture supernatant boiled for 50 min showed no detectable activity and was used as a blank. Product formation was linear with time during the assay, and endochitinase activity was expressed in units of measured absorbance×1000 min⁻¹ according to Wirth and Wolf [21] as a mean of three independent replicates.

Fluorogenic assays using MUF substrate analogues consisted of a sample (20 μl), Na-acetate buffer (75 μl, 50 mM, pH 5.0) and MUF substrate (5 μl, 0.1 mg ml⁻¹). The mixture was incubated in a water bath at 50°C for 30 min, after which the reaction was stopped by addition of Na₂CO₃ (1.4 ml, 0.2 M). The enzyme activity was calculated from the amount of fluorescent MUF product released, as determined with a Perkin-Elmer LS50B fluorometer (excitation and emission wavelengths of 365 nm and 455 nm, respectively; slit size 5.0 nm; Perkin-Elmer Ltd., Buckinghamshire, UK), using pure MUF solution as standard. Product formation was linear during the assay and activities were recorded as a mean of three independent replicates. One unit of enzyme activity was defined as the amount of enzyme releasing 1 μmol of MUF min⁻¹.

2.4 Isoelectric focusing gel electrophoresis

Isoelectric focusing (IEF) gel electrophoresis in combination with a gel overlay reaction with substrate analogues [3] was used to detect the formation of endochitinase isozymes. Samples with equivalent amounts of activity (approximately 10 units of endochitinase activity) were applied to a precast polyacrylamide gel (pH 3.5–9.0; Pharmacia Biotech AB, Uppsala, Sweden). The gel was run according to the manufacturers instructions and was subsequently overlaid with a low temperature-gelling agarose (2% w/v in 0.05 M Na-acetate buffer, pH 5.0) containing CM-chitin-RBV substrate solution (1:1 v/v). The gel was then incubated at 55°C for at least 2 h until clearing bands became apparent. Only major bands showing a distinct clearing were recorded. The isoelectric points (pI) of the clearing bands were determined by comparison with pI markers (Broad pI Calibration Kit; Pharmacia Biotech AB, Uppsala, Sweden) located in the gel by a Coomassie protein stain (PhastGel Blue R, Pharmacia Biotech AB, Uppsala, Sweden). IEF gel electrophoresis was repeated at least once for each isolate.

3 Results

3.1 Chitinolytic activity

All isolates including the two reference strains (CHA0 and BL915) listed in Table 1 were able to hydrolyse the chitin when grown on solid agar medium (10% TSA) supplemented with the chromogenic CM-chitin-RBV substrate (this study and [17]). Clearing zones were observed around the colonies, which suggested that endochitinase activity was excreted from the cells (data not shown).

Fig. 1 shows the results from batch cultures grown to stationary phase in liquid TSB medium. Chitinolytic activities, as defined by the four different substrate analogues, were detected in both the culture supernatant and the cellular fraction. The outgrowth of the cultures was observed within 24 h (data not shown) and the activities recorded after 72 h thus represented late stationary phase. The enzyme activity per unit of cell mass (OD) represented the total, specific activity of enzymes accumulated during logarithmic phase and subsequent stationary phase.

1

Chitinolytic activities of P. fluorescens cultures (stationary phase) grown in Tryptic Soy Broth. Supernatant (black bars) and cell-bound (grey bars) fractions are shown. Enzyme activities were measured using different chromogenic or fluorogenic substrate analogues. Results are presented as enzyme units relative to cell number at sampling time (means of three independent replicates).

The accumulated enzyme activities shown in Fig. 1 showed markedly different patterns among the 12 P. fluorescens isolates, except for the NAGase activities (MUF-N-acetyl-glucosaminide assay), which did not differ significantly. Generally, the two different assays of endochitinase activity (CM-chitin-RBV and MUF-chitotrioside assays) and that of chitobiosidase activity (MUF-chitobioside assay) gave similar results in a given isolate. Isolates expressing high activities with one substrate were also hydrolysing the other substrates at a relatively high rate. The endochitinase activity varied by a factor of approximately 5 between the isolates. Most of the highly productive isolates (BL915, PS7 and 96.578) typically showed both a significant cell-bound activity and an extracellular activity released to the supernatant. In comparison, all the less-productive isolates released all or most of their activity to the supernatant. Finally, all NAGase activity produced by the isolates was also released to the supernatant.

3.2 Growth phase-dependent production of endochitinase activity in selected isolates

In further studies, we tested if there was a growth phase-dependent production of endochitinase activity in three selected isolates, DR54, 96.578 and PS7, representing different P. fluorescens species and biovars (Table 1) and low, intermediate and high levels of endochitinase activity as determined by the CM-chitin-RBV assay (Fig. 1). In this part of the study, we focussed only on the endochitinase activity in the supernatant fraction, since we assumed that this activity would be most relevant for the role of chitinolytic activity in biological control. As shown in Fig. 2, similar growth patterns were observed for the three isolates. The rapid growth for approximately 5 h, followed by slower growth until stationary phase was reached, was characteristic for growth in the complex TSB medium, as has been observed previously during studies of growth phase-dependent production of antibiotic by isolate DR54 (T.H. Nielsen et al. [31]). While endochitinase activity was not detectable during the first phase of rapid growth, all three isolates showed accumulation of enzyme activity during the subsequent phase of slower logarithmic growth, corresponding to 10–25 h of incubation. The results thus indicated that enzyme production was related to cell proliferation during exponential growth, but only shortly while the cultures adopted a slower growth rate in the complex TSB medium. Interestingly, additional enzyme production was observed as the cultures entered stationary phase, corresponding to 25–72 h of incubation. This was less significant in DR54, but was especially evident in isolates 96.578 and PS7. It was also noticed that in all three isolates the accumulated activity was stable up to at least 72 h of incubation, which was the time of sampling for the initial comparisons of total activities per unit of cell mass (Fig. 1).

2

Endochitinase activity (open symbols) and CFU (closed symbols) in P. fluorescens isolates DR54 (A), PS7 (B) and 96.578 (C) during growth in Tryptic Soy Broth. Samples were taken at intervals and results are presented as means of three independent replicates. Standard deviation is shown by error bars.

3.3 Isozyme patterns of endochitinase activity

Isozymic banding patterns were compared to study if the isolate-dependent (Fig. 1) and growth phase-dependent (Fig. 2) production of endochitinase activity (detected as enzyme released to the supernatant) were due to different isozymes. The isozymic patterns of endochitinase activity in the supernatant fraction were chosen for this comparison, but similar results could be obtained using the cellular fraction (data not shown).

Fig. 3 shows the isoelectric focusing pattern of supernatant samples from different growth phases of P. fluorescens DR54. A single isozyme of endochitinase (pI 5.5) was observed during logarithmic growth, while an additional isozyme (pI 5.3) appeared as the culture entered stationary phase after 24 h of incubation. Additional isozymes showing minor bands with faint clearing in the gel could be observed during prolonged incubation (data not shown). A cell-free supernatant sample from a batch culture of this isolate, taken at 15 h of incubation to represent a sample from early logarithmic growth, was incubated as the original batch culture. In this sample both isozyme bands were observed as was the case for the stationary phase culture. The total activity in this cell-free control sample remained constant during the incubation.

3

Isozymic pattern of endochitinase activity produced by P. fluorescens DR54 during growth in Tryptic Soy Broth. Samples were taken at intervals (9 to 96 h) and the supernatant subjected to gel electrophoresis by isoelectric focusing in combination with a substrate overlay technique. Clearing bands represent the different isozymes with endochitinase activity. The gel was scanned (Agfa Foto Look 3.00) and the image was processed by image analysis (Paint Shop Pro 4) to improve contrast.

Table 2 shows that the 12 isolates fell into a total of seven groups, corresponding to different combinations of endochitinase isozymes based on the patterns of major IEF clearing bands: Group A, comprising the high-productive isolate (BL915), had four major bands at low pI values (5.3–5.0 range). Group B, comprising two isolates (DR54 and MN222), had two major bands at low pI values (5.5 and 5.3). Group C, comprising four isolates (DR2, DR3, PS1 and PS21), had two major bands at low pI values (5.4 and 5.6). Group D, comprising another high-productive isolate (PS7), had four major bands at intermediate and low pI values (6.3, 5.8, 5.6 and 5.3). Group E, comprising the isolate 96.578, had five major bands at high, intermediate and low pI values (7.0, 6.5, 6.0, 5.6 and 5.4). Group F, comprising the low-productive isolate MN272, had four major bands at low and intermediate pI values (5.4, 5.6, 6.2 and 6.6). And finally, group G, comprising the third high-productive isolate (MN370), had two major bands at high pI values (6.8 and 7.2). The results clearly demonstrate that patterns of isozyme production between the isolates differ, but that a specific pattern may also be shared by several isolates. Table 2 also shows that a single, native isozyme was found during logarithmic growth phase in each of the different isozyme groups. Group A had this isozyme band at pI 5.3, group B at pI 5.5, group C at pI 5.6, group D at pI 6.3, group E at pI 6.5, group F at pI 6.6 and group G at pI 6.8. Although additional, minor clearing bands appeared for some strains, the major bands noted above were adequate to discriminate the strains into isozyme groupings. Also, the minor bands did not further separate the groupings established by the major bands (data not shown).

2

Endochitinase isozymes in culture supernatant of P. fluorescens isolates during stationary phase when grown in TSB medium

Strain	BL915	DR54	MN222	DR2	DR3	PS1	PS21	CHA0	PS7	96.578	MN272	MN370
Isozyme group	A	B	B	C	C	C	C	C	D	E	F	G
pI
5.0	■
5.1	■
5.2	■
5.3	■a	■	■	■
5.4	■	■	■	■	■	■	■
5.5	■a	■a
5.6	■a	■a	■a	■a	■a	■	■	■
5.7
5.8	■
5.9
6.0	■
6.1
6.2	■
6.3	■a
6.4
6.5	■a
6.6	■a
6.7
6.8	■a
6.9
7.0	■
7.1
7.2	■

^aNative endochitinase isozyme (logarithmic phase).

4 Discussion

4.1 Occurrence of chitinolytic enzymes in P. fluorescens isolates

A complete array of several enzymes comprising chitinolytic activity was demonstrated in the P. fluorescens isolates as based on the definition of enzyme functions by Tronsmo and Harman [3]. Such a complete array of enzymes involved in chitinolytic activity is in accordance with the patterns observed in other bacteria, notably B. licheniformis[22], E. agglomerans[8] and S. marcescens[19]. In the P. fluorescens isolates tested, the chitinolytic activity thus comprised both endochitinase activity, where chitin polymers and longer N-acetyl-glucosamine (GlcNAc) oligomers are hydrolysed, and chitobiosidase activity, where shorter oligomers are hydrolysed into dimer (GlcNAc)₂ units for metabolism within the cell. The isolates also contained a terminal NAGase activity, involved in the final hydrolysis into the monomer (GlcNAc) units. The terminal NAGase activity has sometimes been assigned to chitinolytic activity and even used as an index of chitinolytic activity in both bacterial cultures and environmental samples [23]. In our study, we found a relatively low level of extracellular NAGase activity in all isolates and no detectable activity was ever observed in the cell-bound fraction as expected [22]. These results clearly indicate the absence of a direct relationship between NAGase activity and the other chitinolytic activities in the P. fluorescens isolates. The NAGase activity was therefore not further investigated in this study.

Significant production of chitinolytic activity took place in all isolates when grown on Potato Agar (PA) [17] or in Tryptic Soy Broth (TSB) without exogenous chitin. Although our data show that exogenous chitin is not required in the growth media to obtain production of chitinolytic enzymes, the presence and role of specific inducers of chitinolytic activity in the media obviously cannot be excluded. Production of chitinolytic enzymes in bacteria has thus been reported to be both constitutive [24] and inducible [25]. In this study, we did not perform specific studies to see if inducible, chitinolytic activity could also be found in the P. fluorescens isolates.

Both cell-bound and extracellular (supernatant) components were sometimes observed for the chitinolytic activities. Hence, while all isolates showed extracellular endochitinase activity, some isolates also retained considerable levels of activity in the cell-bound fraction while still others had no activity at all in this fraction. Further, the isolates with endochitinase activity in both cell-bound and extracellular fractions were also the ones showing highest overall activity. Hence, the occurrence of endochitinase activity in the cellular fraction of such high-productive isolates could be due to an intracellular enzyme, but could also result from incomplete release of the enzyme to the extracellular environment.

The quantitative comparison of chitinolytic activities in the P. fluorescens isolates (Fig. 1) showed that endochitinase (CM-chitin-RBV and MUF-chitotrioside assays) and chitobiosidase (MUF-chitobioside assay) activities were interrelated, i.e. similar in terms of activity levels and distribution among isolates. Further, the distribution of endochitinase and chitobiosidase activities between cellular and extracellular fractions was similar among the isolates. The results suggested that the recorded activities either involved coupled enzymes or one single enzyme in the P. fluorescens isolates. It was noticed, however, that endochitinase activity obtained with the CM-chitin-RBV assay showed activities in the cellular fraction in only some of the isolates, while all isolates showed activity in this fraction using the assay with MUF-chitotrioside, the other endochitinase substrate analogue. This indicates that the cell-bound activity in some isolates could hydrolyse longer chitin polymers, while other isolates may only have an exo-N,N′,N″-triacetylchitotriohydrolase activity hydrolysing shorter chitin oligomers. Similar results have been obtained in S. marcescens, where chitobiosidase activity has been observed to combine with either a periplasmic exo-N,N′,N″-triacetylchitotriohydrolase activity or with an extracellular endochitinase activity [19].

In conclusion, the present work shows that several P. fluorescens isolates, showing potential for biological control of plant pathogenic microfungi, contain a complete array of chitinolytic enzymes. The activity was observed in several growth media without exogenous chitin source, and was either extracellular or cell-bound in the cultures. Endochitinase and chitobiosidase activities were tightly coupled and could actually represent the same enzyme. Occasionally, however, the cell-bound fraction appeared to harbour two different endochitinase activities, one hydrolysing CM-chitin-RBV and one hydrolysing MUF-chitotrioside. In an ecological perspective, the constitutive endochitinase activity could be advantageous in biological control since fungal inducers for expression of the antagonistic traits would be unnecessary. Furthermore, constitutive endochitinase activity together with the complete array of chitinolytic enzymes could be a suitable strategy to explore the environment for chitinous material originated from fungi or lower animals in soil. Such a strategy has indeed been proposed for nutrition in a marine Vibrio strain, where the release of N-acetyl-glucosamine oligomers stimulated the production of chitin-binding proteins and probably other chitinolytic enzymes [26]. Since the P. fluorescens isolates originate from barley and sugar beet rhizosphere, they could possibly take advantage from such traits by extending the available substrate sources and thus improving their rhizosphere competence.

4.2 Endochitinase activity and isozymes in P. fluorescens isolates

The appearance of multiple clearing zones in the overlay agar of IEF gels indicated that each of the stationary phase cultures of the P. fluorescens isolates contained multiple isozymes of endochitinase activity as detected with the CM-chitin-RBV substrate analogue. A set of different clearing bands was thus observed in each of the tested isolates and a total of seven banding patterns (Table 2), corresponding to different isozymic groups, could be established. Some banding patterns were unique for one isolate, while other patterns were shared by several isolates. This clearly demonstrated that a number of isozymic groups of endochitinase activity may be found in P. fluorescens. The different IEF gel electrophoretic patterns for endochitinase were not strictly correlated to the biovar affiliation of the P. fluorescens isolates (Table 1), as was the case for other phenetic characters related to fungal antagonism and biological control [17]. The native isozyme produced in logarithmic phase did have the most acidic pI value among isolates of biovar I, however, and the most alkaline pI value was found in isolates of biovar VI (Table 2).

Evidence from the literature indicates that endochitinase production in Pseudomonas spp. grown in batch cultures may show complex regulatory mechanisms as the cultures enter stationary phase and represent high cell densities [14,27]. In the two reference strains, P. fluorescens BL915 and CHA0, production of extracellular enzymes, chitinase [14] and protease [28], was thus shown to be controlled by a global regulatory system characteristic for stationary phase. Interestingly, when three of our own isolates (96.578, PS7 and DR54) were tested for endochitinase production during a full growth cycle of batch cultures (Fig. 2), the overall activity increased during stationary phase for isolates 96.578 and PS7, but not for isolate DR54. This in turn indicated that, in at least some of the P. fluorescens isolates, endochitinase production was under stationary phase regulation [14,29].

The growth experiments further indicated that proteolytic modification of a single, initial endochitinase enzyme produced in logarithmic phase resulted in the multiple isozymes observed as the cultures aged in stationary phase. It may be assumed that the initial enzyme represents the native endochitinase, and that the others appearing at later stages of the growth cultures thus represent either new isozymes being released extracellularly during stationary phase or new forms of the native endochitinase appearing after extracellular proteolysis. It was clear from a control experiment, however, that the additional isozymes appearing during stationary phase in the P. fluorescens cultures could indeed result from proteolytic activity in the supernatants modifying the native endochitinase. It was also noticed that the total activity in this cell-free control sample remained constant, demonstrating that the proteolytic modification of endochitinase did not alter its activity. Such a modification of the native endochitinase was likely to be a cleavage of the protein from one of the terminal ends, without affecting the catalytic activity. Post-translational modification of an endochitinase enzyme by cleavage from the amino-terminal end has been described in B. circulans[5] and B. licheniformis[22], the latter having a total of four endochitinases originating from one native enzyme. Interestingly, a native endochitinase enzyme of Streptomyces olivaceoviridis excreted to the growth medium contained an amino-terminal, proteolytic unit, which was split off in an autocatalytical process releasing the genuine endochitinase [30].

In conclusion, endochitinase activity, in all 12 P. fluorescens isolates, comprised only one native enzyme which was produced primarily in logarithmic phase, but was subsequently modified into several isozymes by a post-translational process in stationary phase. The resulting isozymic patterns grouped the isolates into seven isozymic groups. Production of endochitinase activity seemed in some isolates to be under stationary phase regulation. However, future work should clarify the detailed regulation of gene expression, release and post-translational proteolysis of the endochitinase activity. From the ecological perspective, expression of native endochitinase activity by the P. fluorescens isolates during primary metabolism and cell proliferation could support their efficacy in both biological control and rhizosphere competence. Furthermore, the expression of additional endochitinase activity in at least some P. fluorescens isolates after growth arrest (stationary phase) could possibly be advantageous for maintaining the biological control long after their establishment in the rhizosphere. Before a possible ecological significance of the endochitinase isozymes formed by proteolysis is proposed, further work should clarify if their isoelectric or size characteristics are related to e.g. different affinity towards chitinous substrates or different susceptibility to further proteolysis.

Acknowledgements

This work was supported by the Danish Ministry of Agriculture (contract no. 93S-2466-Å95-00788) in cooperation with Danisco Seed and Novo Nordisk A/S. The reference strains P. fluorescens BL915 and P. fluorescens CHA0 were kindly provided by Dr. Jim Ligon, CIBA Agricultural Biotechnology, USA and Dr. Christoph Keel, University of Lausanne, Switzerland, respectively. The technical assistance of Lene Nielsen and Gitte Larsen is gratefully acknowledged. Finally, we thank Charlotte Thrane for critically reading the manuscript.