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Applied and Environmental Microbiology, August 2000, p. 3632-3636, Vol. 66, No. 8
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
A Simple, Rapid Method for Demonstrating
Bacterial Flagella
Hans-Peter
Grossart,*
Grieg F.
Steward,
Josefina
Martinez,
and
Farooq
Azam
Scripps Institution of Oceanography,
University of California, San Diego, La Jolla, California
92093-0202
Received 13 December 1999/Accepted 24 April 2000
 |
ABSTRACT |
We developed a simple, rapid method for demonstrating flagellation
of bacteria using the fluorescent protein stain NanoOrange (Molecular Probes, Eugene, Oreg.). The NanoOrange reagent binds to
hydrophobic regions of proteins, which results in substantial enhancement of fluorescence. Unbound reagent is essentially
nonfluorescent. NanoOrange fluorescently stained bacterial cell
bodies, as well as flagella and other appendages, which could be
directly observed by epifluorescence microscopy. Detection of flagella
was further improved by using a charge-coupled device camera for image
capture and processing. The reliability of the method was tested by
using 37 pure cultures of marine bacteria. Detection of flagella on the
isolates by NanoOrange staining was compared to detection by
transmission electron microscopy (TEM). For 36 of 37 cultures, the two
methods yielded the same results. In one case, flagella were detected
by TEM but not by NanoOrange, although the difference may be
attributable to differences between the culture preparations. NanoOrange staining is rapid (10 to 15 min) and does not require fixation or dehydration, so live samples can be stained. Since NanoOrange is a general protein stain and works directly in
seawater, it may also prove to be useful for staining other
proteinaceous material that is of interest to aquatic microbial ecologists.
| |
TEXT |
Flagellation is a
useful parameter for identifying certain bacteria, such as
nonfermentative and anaerobic gram-negative bacilli. Clinical studies,
for example, have demonstrated the importance of examining bacterial
flagella for characterization and identification of anaerobic bacteria
(10). Flagellation also appears to be a common
characteristic of bacteria in natural environments. For example,
identification of flagellin gene homologues has been used as a
biomarker in natural communities (21), and in one study, 45 to 70% of the bacteria in coastal seawater samples collected off
Scripps Pier from August to October 1997 were motile (8). The motility exhibited by marine bacteria has been found to be markedly
different from the run and tumble model motility derived from work on
Escherichia coli (16). Visualization of flagella could prove to be useful in studies in which researchers examine the
mechanisms of motility used by diverse bacteria isolated from aquatic environments.
Workers have described a number of methods for staining bacterial
flagella, including the tannic acid-fuchsin method
(10) and modifications of this method
(3, 4, 7, 11-13), as well as silver staining methods
(5, 10, 17, 20). These approaches are satisfactory but
involve complicated protocols. Also, each method has limitations,
such as unstable reagents and fixation-induced alterations. A simple,
rapid method for demonstrating bacterial flagella would facilitate
characterization of clinical and environmental isolates.
In this paper we describe a novel, rapid method for visualizing
bacterial flagella with the fluorescent protein stain
NanoOrange (Molecular Probes, Eugene, Oreg.) and
epifluorescence microscopy. Flagella could be directly observed with a
microscope, but visualization was greatly improved when a
charge-coupled device camera was used for digital image capture and processing.
Bacterial isolates.
Thirty-seven bacterial isolates were
tested (Table 1). These strains had
previously been isolated in the fall and winter of 1993 from seawater
collected ~1 km off Scripps Pier (15). Stock preparations
were kept frozen (
80°C) until they were grown on ZoBell 2216E
medium plates (1.5% [wt/vol] agar [Difco], 5 g of peptone,
and 1 g of yeast extract in 1 liter of prefiltered [pore
size, 0.45 µm] seawater; autoclaved at 121°C for 30 min). To
monitor flagellation during bacterial growth, 50-ml portions of
autoclaved ZoBell 2216E medium were inoculated with bacteria grown on
plates and incubated in 100-ml Erlenmeyer flasks for 26 h at
21°C on a shaker table. Aliquots were taken at 1- to 2-h intervals
during exponential growth and after 23 and 26 h (early stationary
phase). Growth of bacteria was monitored by determining optical density
at 650 nm with a spectrophotometer (Lambda 4; Perkin-Elmer). All values
were corrected with a blank consisting of autoclaved ZoBell 2216E
medium.
Bacterial motility.
Bacterial motility was observed by
dark-field microscopy (16). The equipment used consisted of
a plastic covered depression slide (thickness, 0.5 mm; Fisher) and a
light microscope (Standard 16; Carl Zeiss, Oberkochen, Germany)
equipped with a dry dark-field condenser (0.7/0.85) and a Plan 16/0.35.
Aliquots (100 µl) of culture were transferred onto the depression
slide and observed for 1 min at a magnification of ×320. Bacteria
which exhibited directed movement were considered motile. An isolate
was considered motile even if only a small portion of the cells
displayed motility at the time of sampling.
Detection of bacterial flagella. (i) TEM.
Bacteria were
preserved by adding formaldehyde (final concentration, 2%).
Carbon-stabilized Formvar supports on 200-mesh copper transmission
electron microscopy (TEM) grids (Ted Pella) were rendered hydrophilic
by high-voltage glow discharge (9). In most cases, bacteria
were adsorbed to TEM grids by floating the grids Formvar side down on
an undiluted drop of culture for at least 5 min. In some cases,
aliquots of fixed cultures were diluted to a volume of 10 ml with an
artificial seawater medium and then pelleted onto TEM grids mounted on
acrylic platforms in the bottoms of polyallomer ultracentrifuge tubes
(19). The centrifugation conditions used ranged from 10,000 to 41,000 rpm for 0.5 to 4 h in a swinging bucket rotor (type
SW41; Beckman). The bacteria on grids were stained by submerging the
grids for 20 s in 0.5% (wt/vol) uranyl acetate and then were
rinsed three times (10 s each) by submersion in aliquots of Milli-Q
water. The grids were examined with a Hitachi model H-500 TEM by using
an accelerating voltage of 80 to 100 kV.
(ii) NanoOrange staining.
A 0.5-µl portion of a
NanoOrange stock solution (Molecular Probes) was added to 10 µl of a live bacterial culture on a microscope slide. Samples on the
slide were then mixed with 10 to 20 µl of 30% polyvinylpyrrolidone
(Sigma Chemical Co., St. Louis, Mo.) and covered with a coverslip.
Polyvinylpyrrolidone was used for mounting both to reduce convection
and to serve as a cryoprotectant, which allowed storage of slides for 1 to 2 weeks in a freezer. After waiting 10 to 15 min for staining to
occur, we examined the slide at a magnification of ×1,250 with an
epifluorescence microscope (model BH2; Olympus, Tokyo, Japan) equipped
with a blue filter set (excitation, wavelength, 490 nm; emission
wavelength, 520 nm). The NanoOrange stock preparation was
mainly used undiluted, but a dilution of 1:20 also worked
fine. Higher dilutions (1:100 and 1:1,000) resulted in
reduced image brightness, as judged qualitatively by eye. Staining
times of 10 to 15 min in the dark assured adequate staining, whereas
shorter staining times resulted in images that were qualitatively less
bright. However, prolonged staining times (up to 6 h) did not
change the quality of the image. Images were recorded with a video
camera (model CCD-300 TIFG; Dage MTI) and were examined with an
image-processing system (model IFG-300; Dage MTI) with output to a
black and white video monitor. This system electronically increased
image size by a factor of five and increased the brightness of the
recorded image, which facilitated detection of flagella and other thin
appendages. Images of Escherichia coli (Fig.
1 and 2)
were recorded with a high-speed TEM camera (Mega View II; SIS,
Münster, Germany) mounted on a Zeiss epifluorescence microscope
(Axioskop 2) and were processed by using an image analysis system
(analySIS 3.0; SIS).
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FIG. 1.
Staining of nonmotile, nonflagellated E. coli
cells (a) and motile, flagellated E. coli cells (b and c).
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FIG. 2.
Stalked bacteria on a marine diatom
(Thalassiosira rotula) after NanoOrange staining. Note that
the diatom is strongly stained by the dye, indicating that
large amounts of proteinaceous material were present on the algal
surface.
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|
Bacterial flagella were readily observed by epifluorescence microscopy
after NanoOrange staining and by TEM, as shown for three isolates
in Fig. 3. Scoring for the presence or
absence of flagella by the NanoOrange and TEM methods yielded the
same results for 36 of 37 isolates of marine bacteria grown in a rich medium (Table 1). For strain BBFL6, flagella were observed by TEM but
not by the NanoOrange method. Since different cultures were used to
detect flagella by the NanoOrange method and TEM, the results
obtained for BBFL6 might reflect real differences in flagellation
between the two different preparations rather than a difference between
the techniques. Ideally, the same culture would have been used for both
methods, but the NanoOrange procedure was developed several years
after the more time-consuming TEM work had been completed. Despite the
one equivocal result, our comparison of the two methods indicated that
NanoOrange staining is a reliable method for visualizing flagella
of cultured bacteria.
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FIG. 3.
Comparison of flagellar staining by NanoOrange and
epifluorescence microscopy (a through c) and by TEM (d through f). The
marine bacterial isolates used were SWAT8 (a and d), LHAT4 (b and e),
and LHAT9 (c and f). The organisms in panels a and c are surrounded by
proteinaceous material which is also stained by NanoOrange.
|
|
Detection of flagella with NanoOrange was found to vary with the
growth phase for six cultures of the motile bacteria examined. Flagella were readily observed with the majority of bacteria during exponential growth. In contrast, few or no flagellated cells were observed in the initial lag and late stationary phases. This phenomenon is not unique to the NanoOrange method, since similar results were
obtained with the tannic acid-fuchsin method when other bacteria were
examined in a previous study (14). After 23 to 26 h of incubation, NanoOrange staining also revealed that the cell
surface of most bacteria was covered with proteinaceous material,
which consisted mostly of long fibers or fibrils which may have
represented fimbria, pili, or perhaps some proteinaceous exudate.
Some interesting differences between motility and flagellation
were observed. For example, six isolates (SWAT5, SWAT7, LHFL5, LHAT3, BBFL4, and BB2AT2) were motile as determined by dark-field microscopy, but no flagella were detected by NanoOrange
staining or TEM (Table 1). It is possible that these spiral-shaped
bacteria move by means of periplasmic flagella, as described previously for spirochetes (2). Internal flagella are unlikely to have been detected by either NanoOrange staining or TEM of
unsectioned cells. It is also possible that motility of these strains
is achieved by means other than flagellar rotation. One alternative
mechanism has been proposed for the marine cyanobacterium
Synechococcus sp. (18), which may move by
generating surface waves (6). In another study,
Brahamsha (1) showed that an abundant cell surface
polypeptide is required for swimming by this
nanoflagellated cyanobacterium. Proteinaceous structures
other than flagella were observed on the cell surfaces
of SWAT5, LHFL5, LHAT3, and BBFL4, but whether these structures
are involved in motility was not studied. In addition, two isolates
(SWFL1 and SWAT3) were flagellated but not motile under our
culture conditions. However, one of these isolates (SWAT3) was
motile when it was grown on glucose minimal medium (R. Long, personal
communication), suggesting that motility depended on the type and/or
concentration of substrate.
To prove the validity of our method, we stained the well-characterized
gram-negative bacterium E. coli in its nonmotile,
nonflagellated form as well as in its motile, flagellated form (Fig.
1). Nonmotile cells of E. coli did not exhibit any
flagellation (Fig. 1a), whereas motile cells were surrounded by
numerous flagella (Fig. 1b and c). Unfortunately, our image analysis
system was not capable of resolving single flagella very well, which
made it more difficult to clearly distinguish different types of
flagellation. Since NanoOrange is a general protein stain, it
should also be useful in numerous other applications. For instance,
preliminary results have shown that flagella of protozoans are also
readily stained and visualized (Grossart and Azam, unpublished data).
The fine fibrils observed on bacteria in some stationary-phase cultures suggest that other bacterial appendages, such as the appendages on
stalked bacteria, can be visualized by NanoOrange staining (Fig.
2). The ability to quickly and easily visualize these appendages could
be useful in studies of bacterial attachment to solid surfaces, conjugation, and/or pathogenicity. The fluorescent stain might also be
used in aquatic microbial ecology studies to visualize protein release
by exudation and cell lysis of phytoplankton and bacteria.
In summary, we describe a novel use of the fluorescent protein stain
NanoOrange for rapid visualization of bacterial flagella. Flagella
were discernible by epifluorescence microscopy in exponentially growing
cultures of marine bacteria. Detection of flagella was greatly
facilitated by digital image acquisition with a charge-coupled device
camera and a simple image processor. The NanoOrange method is
simpler and faster than TEM, but it was equally reliable for demonstrating flagellation of a variety of marine bacterial
isolates. Some significant advantages of the NanoOrange
method are that it does not require fixation of the cells, it involves
very little sample manipulation, and staining can be carried out
directly in a growth medium; thus, artifacts due to fixation,
dehydration, and excessive sample manipulation can be avoided.
| |
ACKNOWLEDGMENTS |
We thank Forest Rohwer for discussions.
This work was supported by grants OPP 95-30851 and OPP96-17045 to F.A.
and by grant GR 1540/1-1 (German Science Foundation) to H.-P.G.
| |
FOOTNOTES |
*
Corresponding author. Present address: Institution for
Chemistry and Biology of the Marine Environment, University of
Oldenburg, 26111 Oldenburg, Germany. Phone: 49 441 7980. Fax: 49 441 798 3438. E-mail: hgrossart{at}icbm.uni-oldenburg.de.
Present address: Monterey Bay Aquarium Research Institute, Moss
Landing, CA 95039-0628.
Present address: Department of Microbiology, University of
Barcelona, E-08028 Barcelona, Spain.
| |
REFERENCES |
| 1.
|
Brahamsha, B.
1996.
An abundant cell-surface polypeptide is required for swimming by the nanoflagellated marine cyanobacterium Synechococcus.
Proc. Natl. Acad. Sci. USA
93:6504-6509[Abstract/Free Full Text].
|
| 2.
|
Charon, N. W.,
E. P. Greenberg,
M. B. Koopman, and R. J. Limberger.
1992.
Spirochete chemotaxis, motility, and the structure of the spirochetal periplasmic flagella.
Res. Microbiol.
143:597-603[Medline].
|
| 3.
|
Clark, W. A.
1976.
A simplified Leifson flagella stain.
J. Clin. Microbiol.
3:632-634[Abstract/Free Full Text].
|
| 4.
|
Dowell, V. R., Jr., and T. M. Hawkins.
1974.
Laboratory methods in anaerobic bacteriology: CDC laboratory manual.
Centers for Disease Control, Public Health Service, U.S. Department of Health and Human Services, Atlanta, Ga.
|
| 5.
|
Ederer, G. M., and M. E. Lund.
1981.
Biochemical test procedures, reagents, stains, staining methods, and media, p. 791-838.
In
A. Balows, and W. J. Hausler, Jr. (ed.), Diagnostic procedures for bacterial, mycotic and parasitic infections, 6th ed. American Public Health Association, New York, N.Y.
|
| 6.
|
Ehlers, K. M.,
A. D. T. Samuel,
H. Berg, and R. Montgomery.
1996.
Do cyanobacteria swim using traveling surface waves?
Proc. Natl. Acad. Sci. USA
93:8340-8343[Abstract/Free Full Text].
|
| 7.
|
Forbes, L.
1981.
Rapid flagella stain.
J. Clin. Microbiol.
13:807-809[Abstract/Free Full Text].
|
| 8.
|
Grossart, H.-P., and F. Azam.
1998.
Motility of natural bacterial assemblages in coastal waters.
EOS Trans. (Suppl.)
79:OS27.
|
| 9.
|
Hayat, M. A., and S. E. Miller.
1990.
Negative staining.
McGraw-Hill Publishing Co., New York, N.Y.
|
| 10.
|
Kodaka, H.,
A. Y. Armfield,
G. L. Lombard, and V. R. Dowell, Jr.
1982.
Practical procedure for demonstrating bacterial flagella.
J. Clin. Microbiol.
16:948-952[Abstract/Free Full Text].
|
| 11.
|
Leifson, E.
1930.
A method of staining bacterial flagella and capsules together with a study of the origin of flagella.
J. Bacteriol.
20:203-211[Free Full Text].
|
| 12.
|
Leifson, E.
1938.
Staining of bacterial flagella.
J. Bacteriol.
36:656.
|
| 13.
|
Leifson, E.
1951.
Staining, shape, and arrangement of bacterial flagella.
J. Bacteriol.
62:377-389[Free Full Text].
|
| 14.
|
Leifson, E.
1960.
Atlas of bacterial flagellation.
Academic Press, Inc., New York, N.Y.
|
| 15.
|
Martinez, J.,
D. C. Smith,
G. F. Steward, and F. Azam.
1996.
Variability in ectohydrolytic enzyme activities of pelagic marine bacteria and its significance for substrate processing in the sea.
Aquat. Microb. Ecol.
10:223-230.
|
| 16.
|
Mitchell, J. G.,
L. Pearson,
S. Dillon, and K. Kantalis.
1995.
Natural assemblages of marine bacteria exhibiting high-speed motility and large accelerations.
Appl. Environ. Microbiol.
61:4436-4440[Abstract].
|
| 17.
|
Porter, J. R.,
K. W. Thomulka, and R. A. Smith.
1992.
Demonstrating bacterial flagella.
Am. Biol. Teach.
54:108-111.
|
| 18.
|
Waterbury, J. B.,
J. M. Willey,
D. G. Franks,
F. W. Valois, and S. W. Watson.
1985.
A cyanobacterium capable of swimming motility.
Science
230:74-76[Abstract/Free Full Text].
|
| 19.
|
Wells, M. L., and E. D. Goldberg.
1992.
Marine submicron particles.
Mar. Chem.
40:5-18[CrossRef].
|
| 20.
|
West, M.,
N. M. Burdash, and F. Freimuth.
1977.
Simplified silver-plating stain for flagella.
J. Clin. Microbiol.
6:414-419[Abstract/Free Full Text].
|
| 21.
|
Winstanley, C., and J. A. W. Morgan.
1997.
The bacterial flagellin gene as a biomarker for detection, population genetics and epidemiological analysis.
Microbiology
143:3071-3084[Medline].
|
Applied and Environmental Microbiology, August 2000, p. 3632-3636, Vol. 66, No. 8
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
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