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Applied and Environmental Microbiology, June 1999, p. 2553-2557, Vol. 65, No. 6
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Chitinases from Uncultured Marine
Microorganisms
Matthew T.
Cottrell,
Jessica
A.
Moore, and
David L.
Kirchman*
College of Marine Studies, University of
Delaware, Lewes, Delaware 19958
Received 19 October 1998/Accepted 1 March 1999
 |
ABSTRACT |
Our understanding of the degradation of organic matter will benefit
from a greater appreciation for the genes encoding enzymes involved in
the hydrolysis of biopolymers such as chitin, one of the most abundant
polymers in nature. To isolate representative and abundant chitinase
genes from uncultivated marine bacteria, we constructed libraries of
genomic DNA isolated from coastal and estuarine waters. The libraries
were screened for genes encoding proteins that hydrolyze a fluorogenic
analogue of chitin, 4-methylumbelliferyl 
-D-N,N'-diacetylchitobioside
(MUF-diNAG). The abundance of clones capable of MUF-diNAG hydrolysis
was higher in the library constructed with DNA from the estuary than in
that constructed with DNA from coastal waters, although the abundance
of positive clones was also dependent on the method used to screen the
library. Plaque assays revealed nine MUF-diNAG-positive clones of
75,000 screened for the estuarine sample and two clones of 750,000 for
the coastal sample. A microtiter plate assay revealed approximately 1 positive clone for every 500 clones screened in the coastal library.
The number of clones detected with the plaque assay was consistent with
estimates of the portion of culturable bacteria that degrade chitin.
Our results suggest that culture-dependent methods do not greatly
underestimate the portion of marine bacterial communities capable of
chitin degradation.
| |
INTRODUCTION |
Chitin, a (1
4)--linked
homopolymer of N-acetyl-D-glucosamine, is an
abundant structural polysaccharide produced by many marine organisms.
It is a constituent of the exoskeletons of zooplankton and invertebrate
larvae (10), the cell walls of some chlorophytes (18), and the extracellular material of some diatoms
(3, 28) and prymnesiophytes (5). The first step
in chitin degradation, which is primarily done by microbes
(10), is the hydrolysis of the glycosidic bonds between
N-acetyl-D-glucosamine residues by chitinases
(EC 3.2.1.14). The capacity to degrade chitin is widespread among
taxonomic groups of prokaryotes including the gliding bacteria,
vibrios, Photobacterium spp., enteric bacteria, actinomycetes, bacilli, clostridia (11), and archaea
(12). Bacteria employ several proteins, including
chitin-binding proteins (17, 31), to degrade chitin, but the
hydrolysis by chitinase is the key step in the solubilization and
mineralization of chitin.
The capacity to degrade chitin would seem to be an important attribute
of marine bacteria given the presumed high input of detrital chitin
into the sea (14). Chitinolytic bacteria are typically
detected by either the production of clearing zones on agar containing
chitin or hydrolysis of a fluorogenic substrate analogue of chitin. The
assay for clearing zones suggests that ca. 10% of culturable bacteria
degrade chitin (4, 20, 25), while the portion of strains
hydrolyzing the analogue of chitin has been estimated to be as high as
90% (16). It is not clear which assay produces the more
accurate estimate since both assays have drawbacks; the production of
clearing zones requires export and diffusion of the chitinase into the
surrounding media, while hydrolysis of the analogue may simply reflect
the capacity to degrade small oligomers (2). Furthermore,
whether either culture-based assay reflects the true portion of chitin
degraders in natural bacterial assemblages is unclear, since only a
small fraction (<1%) of the bacteria in seawater can be cultured
(6, 7, 15) and those bacteria in culture are not thought to
be representative of uncultured, natural bacteria (13, 32).
Molecular methods are needed to study chitin degraders without the
isolation of bacteria into pure cultures. Methods that use nucleic acid
probes and PCR primers cannot be designed solely with cultured bacteria
because the nucleotide sequences of chitinase genes from cultured
bacteria so far characterized are very different (33),
suggesting that chitinases from uncultured bacteria may differ greatly
from those in cultured bacteria. Although it is possible that
conservation within groups of chitinases may become clear as more
cultured bacteria are examined, information for cultured bacteria
probably will not be sufficient to design "universal" PCR primers
to retrieve chitinase genes from uncultured bacteria. One alternative
approach that does not rely on conserved nucleotide sequences is to use
genomic libraries to retrieve genes from natural bacterial communities
without cultivation (24, 30, 36).
In this study we used genomic DNA libraries to retrieve chitinase genes
from environmental DNA (26). A high-efficiency lambda phage
cloning vector was used to produce libraries that were screened with a
fluorogenic analogue of chitin to identify chitinase genes. We found
that the frequencies of active clones identified by screening the
libraries by plaque assay were consistent with culture-based estimates
of the portion of marine bacteria that degrade chitin.
| |
MATERIALS AND METHODS |
Sample collection and preparation of DNA.
Coastal seawater
was collected from a depth of 1 m at a station 14 km outside the
entrance to the Delaware Bay estuary in September 1997. Estuarine water
was collected 0.23 km inside the bay in November 1996. Plankton and
particles were collected from the coastal seawater (10 liters) by
filtration onto Gelman Supor filters (0.2 µm) and stored frozen at
80°C in a storage buffer (9). The estuarine sample was
prefiltered through a 0.8-µm (pore-size) polycarbonate filter. Frozen
samples were thawed, and the cells were lysed by using sodium dodecyl
sulfate (SDS) and proteinase K. The lysate was extracted sequentially
with phenol-chloroform and chloroform. RNA was removed by treatment
with RNase A, and the DNA was precipitated with ethanol and further
purified by using the IsoQuick Nucleic Acid Extraction Kit (ORCA
Research, Inc., Bothel, Wash.) according to the manufacturer's instructions.
Size fractionation and cloning of plankton DNA.
The genomic
DNA (5 µg) was prepared for ligation by partial restriction digestion
with the restriction enzyme Tsp509I (New England Biolabs,
Beverly, Mass.) (3.3 U of enzyme per µg of DNA, 65°C, 15 min). The
restriction fragments ranging from 2 to 10 kb were collected by ethanol
precipitation from the 30% portion of a sucrose step gradient (40,000 rpm in a Beckman TLS-55 rotor for 12 h). A 400-ng portion of
restriction fragments was ligated into Lambda Zap II predigested
EcoRI/CIAP-treated vector (Stratagene, La Jolla, Calif.) by
using T4 DNA ligase (Boehringer Mannheim, Indianapolis, Ind.) according
to the manufacturer's protocol. Recombinant lambda phage DNA was
packaged by using Gigapack III packaging extract (Stratagene), and the
titer and fraction of phage containing inserts were determined by
plaque assay with blue-white color selection. The library was amplified
according to the procedure described by the manufacturer of the cloning reagents.
Screening for chitinase genes.
Two approaches were used to
identify clones that hydrolyze the analogue of chitin,
4-methylumbelliferyl
-D-N,N'-diacetylchitobioside (MUF-diNAG). The
first was a plaque assay (4 × 104 plaques per
150-by-15-mm petri plate) screened by spraying MUF-diNAG (50 µM in
100 mM sodium phosphate buffer [pH 8]) onto the plaques as soon as
they were 1 to 3 mm in diameter (22). The library from the
coastal sample was also assayed with the fluorogenic substrate analogue
of cellulose (MUF-cellobioside). Fluorescing plaques were detected by
using a UV (366-nm) light source and transferred to SM buffer (100 mM
NaCl, 0.8 mM MgSO4, 50 mM Tris-HCl, 0.01% gelatin).
Strains of the active phage were purified with two iterations of plaque isolation.
The second approach used an excised copy of the library with each
lambda phage clone represented by a phagemid. The mass excision of the
library was performed by using ExAssist helper phage (Stratagene) according to the manufacturer's protocol to produce phagemids. The
assay was performed in microtiter plates with phagemids adsorbed to
XLOLR cells at a multiplicity of infection of 2 × 10
5. Each well of the plate contained 150 infected cells.
A microtiter plate containing XLOLR cells growing in Luria
broth-tetracycline (12.5 µg/ml) served as a control. After 1 day of
growth, aliquots (75 µl) of the cultures were transferred to sterile
microtiter plates for the MUF-diNAG hydrolysis assay. After the
substrate was added (50 µM), the plates were incubated at 37°C and
examined daily with UV light. Pure strains of positive clones were
obtained from fluorescing wells by two iterations of serial dilution
followed by the isolation of single colonies on agar plates.
Enzymatic activities in protein extracts from positive
clones.
Enzymatic activities of clones were assayed with protein
extracts prepared from recombinant Escherichia coli bearing
plasmids with the cloned DNA. Cells were collected by
centrifugation, washed three times with Tris-buffered saline (20 mM
Tris, 150 mM NaCl [pH 7.5]), resuspended in Tris-buffered saline, and
sonicated. Sarcosyl (1% [wt/vol]) was added to the lysate before
incubation on ice for 1 h. Particulate matter was removed from the
extract by centrifugation (9,000 × g, 10 min), and the
activities of the supernatant were assayed.
The capacity of the protein extracts to hydrolyze the fluorogenic
substrate analogues 4-methylumbelliferyl
N-acetyl-
-
D-glucosaminide
(MUF-NAG),
MUF-diNAG, and 4-methylumbelliferyl
-
D-
N,
N',
N"-triacetylchitotrioside
(MUF-triNAG) was determined with 50 µM substrate additions and
incubation at 37°C. At time intervals ranging from a few minutes
to
an hour, subsamples were removed and added to glycine carbonate
buffer
(pH 9.7) in order to measure MUF
fluorescence.
The hydrolysis of glycol chitin by the protein extracts was assayed by
glycol chitin-SDS-polyacrylamide gel electrophoresis
(
33).
Electrophoresis was performed with an 8% polyacrylamide
gel containing
0.1% SDS and 0.01% glycol chitin. After electrophoresis,
the gel was
incubated at 37°C for 2 h in 100 mM sodium acetate
buffer (pH
5.0) containing 1% Triton X-100. The gel was stained
for 5 min in
0.01% Calcofluor White M2R in 0.5 M Tris-HCl (pH
9.0) and destained
overnight in water. Equal amounts of protein
(200 µg) were
electrophoresed without prior heat treatment. Clearing
zones produced
by the hydrolytic activity of the extracts were
visualized with UV
light (366
nm).
| |
RESULTS |
Numbers of cloned chitinase genes recovered from libraries.
The coastal library was screened by using the fluorogenic analogues of
chitin (MUF-diNAG) and cellulose (MUF-cellobiose) to detect chitinase
and cellulase genes, respectively. Two MUF-diNAG-hydrolyzing clones
designated pG1 and pG2 were isolated after 7.5 × 105
clones were screened by the plaque assay. Restriction digestion of
clones pG1 and pG2 with a mixture of XbaI and
KpnI produced identical patterns composed of 1.4- and 4-kb
bands plus a 2.9-kb band representing the pBluescript KS() vector,
indicating that these clones have identical 5.4-kb inserts (Fig.
1). Digestion with the four-base cutter
Tsp509I produced identical restriction patterns (data not
shown). Cellulases were not detected since no plaques fluoresced after
application of MUF-cellobioside in the plaque assays.
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FIG. 1.
Restriction patterns (XbaI with
KpnI) of clones that hydrolyze MUF-diNAG. (A) Clone
identified by screening the library by plaque assay. Lanes: 1, pG1; M,
molecular weight marker. (B) Clones identified by screening the library
in microtiter plates. Lanes: 1, p2F9; 2, p3A6; 3, p4H11; 4, p5C7; 5, p5F2; 6, p6D5; 7, p6E11; 8, p7C8; 9, p7D9; 10, p7F6; 11, p9D5; 12, p10D3; 13, p14E11; M, molecular weight marker.
|
|
Screening a total of 2.3 × 10
5 clones from the
coastal library in microtiter plates with MUF-diNAG yielded 432 fluorescing
wells. Clones from 14 fluorescing wells were purified by
dilution
series and spreading on agar plates. Thirteen wells remained
MUF-diNAG
positive after purification to single colonies. Restriction
digestion
with a mixture of
XbaI and
KpnI
revealed that all 13 were different
and were not the same as the clones
isolated by the plaque assay
(Fig.
1). The size of the cloned inserts,
estimated by summing
the restriction fragments, ranged from 1.8 to 4.2
kb.
The estuarine library was screened for chitinases by the plaque assay
with MUF-diNAG. Nine clones hydrolyzing MUF-diNAG were
identified after
75,000 clones were screened (see Table
2). Restriction
digestion with
EcoRI revealed insert sizes ranging from 5.0 to
6.1 kb (data
not
shown).
Clone phenotypes.
The phenotypes of 13 clones from the coastal
library were characterized, and the enzymes they produced were
classified by assaying protein extracts for hydrolysis of various
fluorogenic N-acetylglucosamine oligomers (19).
Four clones (p2F9, p3A6, p4H11, and p14E11) hydrolyzed all three
N-acetylglucosamine oligomers, suggesting that they produce
exochitinases (Table 1). The enzymes produced by clones p5F2, p6E11, and p10D3 were classified as
chitobiosidases because they hydrolyzed only MUF-diNAG. Clone p5C7
hydrolyzed MUF-NAG and MUF-diNAG, suggesting that it might produce an
exochitinase; however, it did not hydrolyze MUF-triNAG (Table 1).
Protein extracts prepared from five clones had no activity against any
of the MUF substrates, even though cultures of
E. coli bearing these plasmids hydrolyzed MUF-diNAG at rates 7- to 34-fold
higher than control cells possessing the pBluescript KS(
) vector
(Table
1). The activity of protein extracts from all clones identified
with the microtiter plates was less than that of the intact cells
(Table
1) and decreased upon lysis of the cells by either sonication
or
lysozyme.
The estuarine library contained clones pJAM6 and pJAM9 that hydrolyzed
all three chitin analogs, and the enzymes they produced
were classified
as exochitinases (Table
2). Clone pJAM19
was
active against MUF-diNAG and MUF-triNAG but not MUF-NAG, suggesting
it produced an endochitinase. Because clones pJAM4 and pJAM5 were
active against only MUF-diNAG, the enzymes they produce were classified
as chitobiosidases (Table
2). The relative rates of hydrolysis
of the
various analogues varied among the clones. Clone pJAM4
and clone pJAM5
had the highest activities (10- to 50-fold above
background) against
MUF-diNAG. The remaining clones hydrolyzed
the various analogues at
rates 2- to 10-fold above background.
Protein extracts were assayed for the capacity to hydrolyze glycol
chitin in polyacrylamide gels containing this soluble form
of chitin.
After electrophoresis, the gels were stained with Calcofluor
to
visualize areas of the gel in which chitin had been hydrolyzed.
Four
clones from the coastal library (p2F9, p3A6, p4H11, and p5C7)
made
clearing zones that were distinguishable from the control
[pBluescript
KS(
)] (Fig.
2). The enzymes appeared
to have molecular
masses of 98 and 250 kDa. However, their actual sizes
cannot be
measured from this analysis, as proteins do not run true to
their
molecular masses in this type of gel (
34). Protein
extracts
of four clones (pJAM5, pJAM6, pJAM9, and pJAM19) from the
estuarine
library hydrolyzed glycol chitin after SDS-electrophoresis.
Clone
pJAM5 produced an active enzyme that appeared to have a molecular
mass of 250 kDa. Clones pJAM9 and pJAM19 produced a clearing at
ca. 98 kDa. Clone pJAM6 produced many regions of clearing between
250 and 30 kDa (data not shown).
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FIG. 2.
Calcofluor-stained glycol chitin gel of proteins
extracted from E. coli bearing the plasmids pBluescript
KS() (lane 1), p2F9 (lane 2), p3A6 (lane 3), p4H11 (lane 4), and p5C7
(lane 5). Regions of the gel with hydrolyzed chitin were
distinguishable by their lack of fluorescence (dark bands).
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|
| |
DISCUSSION |
The screening of genomic DNA libraries for the expression of
targeted genes has been used to collect genes encoding proteins with
industrial applications (26), but this approach can be used
to ask ecological questions as well. In this study, we cloned genes
coding for proteins that hydrolyze a fluorogenic analogue of chitin and
glycol chitin from environmental DNA. Examining chitinase genes is
important for understanding the ecology of chitin-degrading bacteria
and chitin degradation in aquatic systems.
Screening environmental DNA libraries has its limitations. Detection of
a cloned chitinase by expression requires cloning a native promoter
with the chitinase gene or the alignment of the cloned gene with the
reading frame of the lacZ promoter on the vector
(22). In some cases, the inability to obtain an expressed clone is not clear. For example, part of the chiA gene of
Vibrio harveyi was obtained by screening a plasmid library
for activity (29), but a full-length chiA was
never obtained in a library made in lambda phage (34).
Furthermore, the fidelity with which a genomic library reflects the
abundance of genes in the community can be further influenced by
manipulations of the library itself. For example, clones pG1 and pG2,
which have the same restriction fragment length polymorphism pattern,
could represent genes from two separate genomes captured by the
library; alternatively, one may be a duplicate produced when the
library was amplified to create copies of the library. It is possible
to screen a library without amplification, but copies of the library
were necessary in order to screen with more than one substrate analogue.
In spite of the limitations, genomic DNA libraries screened for gene
expression are valuable tools for ecological studies because they can
provide information about enzymes from organisms without cultivation.
Furthermore, they are well suited for genes such as those encoding
chitinases which do not have regions of similarity needed for designing
universal probes or PCR primers. In addition to recovering previously
unrecognized diversity, eventually we wish to build molecular
approaches to examine the frequency and expression of chitinases in
natural samples and to determine the relative abundance of organisms
that possess chitinases. Answering these questions will require
extensive development of methods to detect genes in natural
bacterial assemblages. Although a direct estimate of chitinolytic
bacteria is not technically possible now, we can use our data to obtain
an indirect estimate of how many bacteria degrade chitin. In addition
to being ecologically interesting, the calculation is important for
determining if the frequency of MUF-diNAG-positive clones we obtained
is reasonable.
We used the abundance of clones hydrolyzing MUF-diNAG to estimate the
portion of uncultured bacteria that are chitinolytic. The estimation
was based on the relationship among insert size, genome size, and the
number of clones that must be screened to find a single-copy gene in a
library with a probability of 0.99 (1):
![N = <UP>ln</UP>(1 − 0.99)/<UP>ln</UP> [1 − (<UP>insert size/genome size</UP>)]](/content/vol65/issue6/fulltext/2553/img001.gif)
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(1)
|
where
N is the number of clones that must be screened.
The calculation was made by using a genome size representative of
bacteria (2 × 10
6 kb) and an insert size of 4 kb.
Because the coastal library was
constructed with total plankton DNA,
which typically contains
about 50% bacterial DNA (
21,
38),
the number of clones to
screen for this library was multiplied by
2.
We expected more MUF-diNAG-positive clones for a given library size,
and thus a higher frequency of positive clones than indicated by
equation 1, because bacteria typically have not one but about five
chitinase genes. Thus, the expected frequency of MUF-diNAG positive
clones is:
|
(2)
|
where
N is calculated from equation 1. We assumed five
chitinase genes per chitinolytic species based on observations of
the
enzymes produced by five chitin-degrading bacteria.
Serratia marcescens (
8) and
Streptomyces
olivaceoviridis (
23) each
produce five chitinases,
while
Streptomyces plicatus (
22) and
Bacillus circulans (
37) produce four and six
chitinases, respectively.
It was assumed that the number of chitinase
genes per bacterium
is equal to the number of chitinases produced. The
number of chitinase
enzymes may be larger than the number of chitinase
genes because
chitinases may be modified by proteolytic cleavage to
produce
additional chitinases (
23,
35). For example,
V. harveyi appears
to produce 10 chitinases by proteolytic
cleavage of proteins encoded
by five chitinase genes (
34).
The type and number of chitinases
produced varies with exposure to
different types of chitin (
34),
so estimates of the numbers
of chitinases produced by various
bacteria may increase as more types
of chitin are tested, but
probably by no more than a factor of
2.
The expected number of MUF-diNAG-positive clones is a maximum estimate
because our calculation so far assumes that all genomes represented in
the library contain chitinase genes. Thus, the percentage of
chitin-degrading bacteria in the original water sample can be estimated
by dividing the observed frequency of MUF-diNAG positive clones by the
expected frequency. The estimates of the percentages of chitin
degraders for the estuarine and coastal communities were 5.5 and
0.12%, respectively, based on the results from plaque assays (Table
3). In contrast, screening the coastal library as phagemids in microtiter plates produced far more
MUF-diNAG-positive clones than would be expected if all of the bacteria
in the community were chitin degraders. The explanation for this large
number of MUF-diNAG-positive clones is not clear, but several of these
clones probably produce proteins that hydrolyze the substrate analogue but are not involved in the hydrolysis of chitin.
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TABLE 3.
Percentage of chitinolytic bacteria in coastal and
estuarine samples inferred from the frequency of clones
hydrolyzing MUF-diNAG
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|
The microtiter assay seems to detect low-level hydrolysis of MUF-diNAG
not observed with the plaque assay. This greater sensitivity is likely
because a microtiter well contains more cells than a plaque and because
the microtiter assay uses intact cells containing excised plasmids,
whereas cells in plaques have been lysed. In addition to likely higher
enzyme production, enzyme activity was probably also higher in the
microtiter assay with intact cells since we found that lysing
MUF-diNAG-positive clones by sonification, which is analogous to viral
lysis, greatly reduced the hydrolysis of MUF-diNAG. Finally, diffusion
of the fluorescent signal away from plaques may be another contributing
factor. Robbins et al. (22) suggested that a low signal can
limit the detection of clones hydrolyzing a fluorogenic substrate and
pointed out that the accumulation of fluorescence requires the
production of the fluorescing compound to exceed its removal by
diffusion. The microtiter wells were not subjected to this limitation
and fluorescence was maintained for several days.
Given the prevalence of chitin-producing organisms in the sea, it might
be anticipated that most marine bacteria are able to degrade chitin.
Studies with cultures, however, suggest that relatively few marine
bacteria degrade chitin, ranging from 0.4 to 19% of total cultured
bacteria (4, 20, 25). Our estimates for estuarine and
coastal waters were within this range. Although chitinase activity can
be much higher on particles than in the surrounding seawater
(27), including particle-associated bacteria in the coastal
library did not greatly increase the estimate of the portion of
bacteria that degrade chitin. Even though culture-based methods
retrieve only a small fraction of the total bacteria, our results
suggest that culture-based estimates of the percentage of chitinolytic
bacteria may be correct. However, there still remains a large pool of
uncultured chitin-degrading bacteria in aquatic environments, and
describing their chitinases will produce a better understanding of
chitin degradation in the sea.
| |
ACKNOWLEDGMENTS |
This research was funded by the U.S. Department of Energy. The
collection of samples was made possible by a National Science Foundation grant.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: College of
Marine Studies, University of Delaware, 700 Pilottown Rd., Lewes, DE
19958. Phone: (302) 645-4375. Fax: (302) 645-4028. E-mail:
kirchman{at}udel.edu.
| |
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Abstract
Introduction
