Download Canadian Journal of Microbiology 47

1
Ultrastructure of interaction in alginate beads
between the microalga Chlorella vulgaris with its
natural associative bacterium Phyllobacterium
myrsinacearum and with the plant growthpromoting bacterium Azospirillum brasilense
Vladimir K. Lebsky, Luz E. Gonzalez-Bashan, and Yoav Bashan
Abstract: Chlorella vulgaris, a microalga often used in wastewater treatment, was coimmobilized and coincubated either with the plant growth-promoting bacterium Azospirillum brasilense, or with its natural associative bacterium
Phyllobacterium myrsinacearum, in alginate beads designed for advanced wastewater treatment. Interactions between
the microalga and each of the bacterial species were followed using transmission electron microscopy for 10 days. Initially, most of the small cavities within the beads were colonized by microcolonies of only one microorganism, regardless of the bacterial species cocultured with the microalga. Subsequently, the bacterial and microalgal microcolonies
merged to form large, mixed colonies within the cavities. At this stage, the effect of bacterial association with the
microalga differed depending on the bacterium present. Though the microalga entered a senescence phase in the presence of P. myrsinacearum, it remained in a growth phase in the presence of A. brasilense. This study suggests that
there are commensal interactions between the microalga and the two plant associative bacteria, and that with time the
bacterial species determined whether the outcome for the microalga is senescence or continuous multiplication.
Key words: Azospirillum, Chlorella, Phyllobacterium, wastewater treatment, water bioremediation.
Résumé : Chlorella vulgaris, une algue microscopique fréquemment utilisée dans le traitement des eaux usées, a été
co-immobilisée et co-incubée avec soit la bactérie Azospirillum brasiliense favorisant la croissance végétale, soit la bactérie Phyllobacterium myrsinacearum avec qui elle est naturellement associée, dans des billes d’alginate conçues pour
l’épuration avancée des eaux usées. Les interactions entre les algues microscopiques et chacune des espèces de bactéries furent suivies pendant 10 jours par microscopie électronique à transmission. Au début, la plupart des petites cavités
à l’intérieur des billes étaient colonisées par des microcolonies de seulement un micro-organisme, sans égard à l’espèce
bactérienne co-cultivée avec l’algue. Par la suite, les colonies de bactéries et d’algues microscopiques fusionnèrent pour
former de grosse colonies mixtes à l’intérieur des cavités. À ce stade, l’effet de l’association entre la bactérie avec
l’algue variait selon le type de bactérie présente. L’algue demeura dans sa phase de croissance en présence de A. brasiliense, alors qu’elle entra dans une phase de sénescence en présence de P. myrsinacearum. Cette étude indique qu’il
existe des interactions commensales entre l’algue microscopique et deux bactéries s’associant aux plantes, et que la période de contact avec l’espèce bactérienne détermine si le résultat sera la sénescence ou une multiplication continuelle.
Mots clés : Azospirillum, Chlorella, Phyllobacterium, traitement des eaux usées, bioremédiation des eaux.
[Traduit par la Rédaction]
Lebsky et al.
8
Introduction
Over the last two decades, microalgae and bacteria have
been immobilized in a variety of polymers, the most common being alcjmginate of various forms and shapes. Microbes contained within polymers provide a convenient
inoculum for numerous industrial, environmental, and agri-
cultural applications (Bashan 1998; Cassidy et al. 1996;
Fages 1992; Leenen et al. 1996; Omar 1993; Tanaka and
Nakajima 1990). For example, the microalga Chlorella sp.
coimmobilized in a polymer with other microorganisms has
been used in biotechnological processes where the microalga
provides oxygen for the accompanying microorganism involved in the compound transformation (Chevalier and De la
Received March 29, 2000. Revision received September 14, 2000. Accepted September 18, 2000. Published on the NRC Research
Press web site on December 12, 2000.
V.K. Lebsky and Y. Bashan.1 The Center for Biological research of the Northwest (CIB), P.O. Box 128, La Paz, B.C.S. 23000,
Mexico.
L.E. Gonzalez-Bashan. The Center for Biological research of the Northwest (CIB), P.O. Box 128, La Paz, B.C.S. 23000, Mexico,
and Department of Biology, Pontificia Universidad Javeriana, Apartado Aereo 56710, Santafe de Bogota, Colombia.
1
Author to whom all correspondence should be addressed (e-mail: [email protected]).
Can. J. Microbiol. 47: 1–8 (2001)
DOI:10.1139-cjm-47-1-1
© 2001 NRC Canada
2
Noüe 1988; Khang et al. 1988; O’Reilly and Scott 1995).
When immobilized in alginate beads and applied experimentally in semiarid agriculture (Bashan 1998; Bashan et al.
1987), the bacterium A. brasilense has been found to survive
for prolonged periods (Bashan and Gonzalez 1999). As a
seed or root inoculant, diazotrophic A. brasilense nonspecifically promotes the growth of many terrestrial plants,
and increases the yield of numerous crop plants (Bashan and
Holguin 1997a, 1997b; Okon and Labandera-Gonzalez
1994). It is thus known as a plant growth-promoting bacteria
or PGPB (Bashan and Holguin 1998).
Phyllobacterium myrsinacearum is found in leaf nodules
on a few tropical plants (Knösel 1962, 1984) and in the
rhizoplane of sugar beet (Lambert et al. 1990). Recently, we
isolated a strain of P. myrsinacearum from a culture of the
microalga C. vulgaris obtained from a wastewater-stabilization
pond (Gonzalez et al. 1997), where it maintained a close association with the microalga (Gonzalez-Bashan et al. 2000).
Chlorella vulgaris, a fresh water microalga (Oh-Hama and
Miyachi 1992), is often used for tertiary wastewater treatment (Gonzalez et al. 1997; De la Noüe and Pauw 1988; Lau
et al. 1997; Tam et al. 1994) and for several industrial processes (Kayano et al. 1981; Wikstrom et al. 1982).
Although numerous bacteria are known to influence the
growth and development of higher plants, the response of
microalgae to artificial bacterial inoculation is, to the best of
our knowledge, unknown. Two recent studies showed that
growth of C. vulgaris was enhanced following inoculation
with A. brasilense, but not in response to inoculation with its
natural associative bacterium P. myrsinacearum. Both bacterial inoculants significantly increased pigment production of
the microalga, and changed its nitrogen and phosphorus metabolism (Gonzalez and Bashan 2000; Gonzalez-Bashan et
al. 2000).
The aim of this study was to further characterize, at the
ultrastructural level, the response of C. vulgaris to bacterial
inoculation. Chlorella vulgaris was coimmobilized separately with A. brasilense and P. myrsinacearum, and the pattern of colonization of the internal cavities of the alginate
beads was followed for ten days by transmission electron
microscopy. A model of the possible interactions among
microalga, bacterium, and the polymer matrix is presented
and discussed.
Materials and methods
Microorganisms and axenic growth conditions
Chlorella vulgaris Beijerinck, (UTEX 2714, U.S.A.) isolated
from the secondary effluent of a wastewater-treatment stabilization
pond near Santafe de Bogota, Colombia (Gonzalez et al. 1997),
was purified from associative bacteria using a mixture of antibiotics (Gonzalez-Bashan et al. 2000). Prior to immobilization in
alginate beads, axenic C. vulgaris was cultivated in sterile mineral
medium (C30) (Gonzalez et al. 1997) for 5 days. Azospirillum
brasilense Cd (DSM 7030, Germany) was grown in liquid Nutrient
Broth (Oxide, U.K.) or N-free OAB medium at 30 ± 2°C for 16 h
using standard methods (Bashan et al. 1993). For viscosity measurements of solutions containing alginate, A. brasilense was
grown as above in nutrient broth medium supplemented with 1%
alginate, at a viscosity described below for 5 days. Viscosity of the
growth medium was measured by viscosimeter (Brookfield,
LVTDV-I, U.S.A.) (6 rpm, 29°C). Phyllobacterium myrsinacearum
Can. J. Microbiol. Vol. 47, 2001
BOG-1–98 was grown in liquid nutrient broth (Oxide, U.K.) at
30 ± 2°C for 16 h.
Coimmobilization of Chlorella vulgaris and Azospirillum
brasilense or C. vulgaris and Phyllobacterium
myrsinacearum in alginate beads
Pure cultures of microorganisms were immobilized in alginate
beads using the method described by Bashan (1986). Briefly,
20 mL of axenically grown cultures of C. vulgaris containing approximately 6.0 × 106 cells/mL were mixed with 80 mL of sterile,
6000 centipois (cps), 2% alginate solution (a solution made from
14 000 and 3500 cps alginate, Sigma) and stirred for 15 min. The
solution was dripped from a sterile syringe into 2% CaCl2 solution
(Bettmann and Rehm 1984) under slow stirring. The beads formed
were left for 1 h at 25 ± 2°C for curing and then washed in sterile
saline solution (0.85% NaCl). Where cocultures of either
A. brasilense or P. myrsinacearum and the microalga were coimmobilized (separately), the same concentration of microalga as
used for immobilization of the pure culture was mixed with each of
the bacterial species (approximately 109 cfu/mL) prior to bead formation, but the volume of each microbial culture was reduced to
10 mL before adding the alginate. Because immobilization normally reduces the number of microorganisms in the beads (Bashan
1986), a second overnight incubation step was necessary as described below.
Culture conditions for coimmobilized microorganisms
Coimmobilized microorganisms were grown in batch cultures in
the mineral salts of residual water medium (RWM)(Gonzalez et al.
1997) containing the following (mg/L): NaCl, 7; CaCl2, 4;
MgSO4·7 H2O, 2; K2HPO4, 21.7; KH2PO4, 8.5; Na2HPO4, 33.4;
NH4Cl, 10. The level of phosphate in the medium was insufficient
to dissolve the constructed beads. Cultures (500 mL) were incubated in Erlenmeyer flasks at 25 ± 2°C, 150 rpm, and with a light
intensity of 60 µmol·m–2·s–1 for 10 days. Samples for analysis were
taken aseptically.
Transmission electron microscopy
Intact beads were first fixed with 2.5% glutaraldehyde in 0.1 M
cacodylate buffer (pH 7.2) for 2 h at 20 ± 2°C, rinsed, and then
postfixed in 1% osmium-tetraoxide in the same buffer for 1 h at the
same temperature. The beads were then rinsed with distilled water
and dehydrated in a series of increasing ethanol concentrations
(30–100%) for 20 min each, and finally with 100% acetone. The
dehydrated beads were embedded in Araldite resin (Sigma).
Ultrathin sections were cut using a Reichert-Young Ultracut
ultramicrotome (Vienna, Austria), stained with uranyl acetate and
lead citrate, and visualized using a Hitachi H-300 transmission
electron microscope (Japan) at 70 kV. Because each bead contained only two microorganisms (microalga and one bacterial species) at the time of coimmobilization, and all procedures were done
under aseptic conditions, immunolabelling of A. brasilense or
P. myrsinacearum cells was not required for later identification in
the photomicrographs.
Experimental design and sampling
Cultures were prepared in triplicate (one flask is a single replicate) and each experiment was repeated twice. Controls were prepared similarly but without microorganisms in the beads. Five
beads were taken randomly from each flask for electron microscopic analysis. Over 300 photomicrographs were taken during this
study and a representative sample is presented. Viscosity measurements were done in triplicate (one flask is a single replicate).
© 2001 NRC Canada
Lebsky et al.
3
Fig. 1. Immobilization of the wastewater-treatment microalga C. vulgaris in alginate bead. (a) C. vulgaris cell immediately after immobilization filling the entire small cavity in a bead. (b) Three C. vulgaris cells within a cavity 2 days after inoculation. (c) Two
C. vulgaris cells and a ghost cell of C. vulgaris within the cavity. (d) Aging C. vulgaris cells after 10 days. Bars represent 1 µm. Abbreviations: Al – alginate bead; c – cavity inside the bead; ch – C. vulgaris; chg – ghost cell of C. vulgaris; EC – electron-dense material of unidentified nature; g – grana; l – cytoplasmic lipids; N – nucleus; St – starch granules.
Results and discussion
Colonization of the roots of numerous plant species by
Azospirillum sp. has been well documented using a large variety of visual techniques including electron microscope
(Assmus et al. 1995; Bashan et al. 1986, 1991; Bashan and
Levanony 1989; Berg et al. 1979; Levanony et al. 1989;
Puente et al. 1999; Schank et al. 1979; Vande Broek et al.
1993). Only a single scanning-electron- microscope study
shows formation of A. brasilense microcolonies on the surface of alginate beads following immobilization (Bashan
1986). To the best of our knowledge, no photomicrographs
are available to illustrate the ultrastructure of C. vulgaris or
P. myrsinacearum immobilized in synthetic beads.
By transmission electron microscopy, we followed events
occurring over a period of 10 days in the interior of the
alginate beads colonized by the microalga C. vulgaris
coimmobilized and cocultured with either its natural associative bacterium P. myrsinacearum or with the common plant
growth-promoting bacterium A. brasilense. These bacteria
have previously been shown to affect C. vulgaris metabolism
(P. myrsinacearum) and growth rate (A. brasilense) (Gonzalez and Bashan 2000; Gonzalez-Bashan et al. 2000).
Though the events following coimmobilization and
coculturing of C. vulgaris with each of the two bacterial species appeared similar initially, the outcome after ten days
was different. Regardless of the partner bacterial species,
immediately upon coimmobilization, cells of the microalga
and bacterium were uniformly trapped within the alginate
matrix in cavities formed by the solidification process. Similar cavities were formed in the absence of microorganisms in
control beads. During the initial coculturing period (24 h after coimmobilization), small, uniform colonies formed
within the alginate beads in cavities of various sizes (several
µm to 0.3 mm) (Figs. 1a, 2a, 3a, 4 Stage 1). The cavities
contained either microalgal or bacterial cells, but not both.
As found previously (Gonzalez and Bashan 2000; GonzalezBashan et al. 2000), during the first five days of incubation,
all microorganisms multiplied inside the beads.
Individual bacterial colonies apparently moved close to
the immobile microalgal colonies (Figs. 2b, 3b, 4 Stage II).
Although we do not have ultrastructural evidence of how
this movement occurred, some explanations are feasible. It is
known that upon formation, the alginate beads contained a
large number of cavities, some of which are connected
(Bashan 1986). The bacteria might migrate internally along
© 2001 NRC Canada
4
Can. J. Microbiol. Vol. 47, 2001
Fig. 2. Coimmobilization of the wastewater-treatment microalga C. vulgaris and its associative bacterium P. myrsinacearum in alginate
beads. (a) C. vulgaris and P. myrsinacearum cells occupying separate cavities immediately after immobilization. (b) C. vulgaris cell
approached by P. myrsinacearum colony. (c) Dividing (arrows) P. myrsinacearum cell in the bead cavity. (d) C. vulgaris and
P. myrsinacearum cells sharing the same bead cavity. (e) Dividing (arrows) C. vulgaris cell in an aging culture containing ghost cells
after 10 days. (f) C. vulgaris cell and a ghost cell of C. vulgaris sharing the cavity with P. myrsinacearum cells in an aging coculture.
(g) Aging C. vulgaris and P. myrsinacearum cells after 10 days where most cells are ghost cells. Bars represent 1 µm. Abbreviations:
Al – alginate bead; c – cavity inside the bead; ch – C. vulgaris; chg – ghost cell of C. vulgaris; EC – electron-dense material of unidentified nature; l – cytoplasmic lipids; p – pyrenoid; Ph – P. myrsinacearum; Phg – P. myrsinacearum ghost cell.
the formed polymer chains of the beads (Mchugh 1987).
Both A. brasilense and P. myrsinacearum are motile bacteria
and may be responding chemotactically to signals from the
microalga, such as those thought to be present in exudates of
higher plants (Bashan and Holguin 1994; Knösel 1984). Alternately, the bacteria may partially dissolve the alginate,
© 2001 NRC Canada
Lebsky et al.
5
Fig. 3. Coimmobilization of the wastewater-treatment microalga C. vulgaris and the plant growth-promoting bacterium A. brasilense in
alginate beads. (a) C. vulgaris and A. brasilense cells occupying separate cavities immediately after immobilization. (b) A C. vulgaris
cell approached by a A. brasilense colony. (c) Enlarged view of C. vulgaris with A. brasilense cells showing an electron dense material
separating the two microbial species within the bead cavity. (d) C. vulgaris and A. brasilense cells sharing the same bead cavity.
(e) C. vulgaris and A. brasilense cells sharing a bead cavity in an aging coculture after 10 days. Note the absence of ghost cells of
both microorganisms as seen in Fig. 2. Bars represent 1 µm. Abbreviations: Al – alginate bead; Az – A. brasilense; c – cavity inside
the bead; ch – C. vulgaris; EC – electron-dense material of unidentified nature; St – starch granules.
thereby removing obstacles to movement. The capacity to
degrade alginate is known in soil bacteria (Dyrset et al.
1994) and marine bacteria (Uchida and Nakayama 1993),
but has not yet been confirmed for P. myrsinacearum. However, Azospirillum is known to produce small quantities of
acids, like indole-3-acetic acid and others (Baca et al. 1994)
that might dissolve the thin alginate layers (0.3 µm or less)
initially separating the microalgal and the bacterial colonies.
Viscosity measurements taken in this study of A. brasilense
culture medium supplemented with alginate showed a decrease in viscosity of 9–15%. Finally, the multiplication of a
large population of each microorganism inside the beads
may create an internal force fracturing the beads and allowing movement along the fractures.
After contact, small colonies merged to form one larger
colony composed of a mixture of the abundant bacterial and
algal cells that filled the cavity (Figs. 2d, 3d). At the same
time, the walls of the external cavities at the periphery of the
bead began to deteriorate forming large open cavities
(Fig. 4, stages II and III). That alginate deterioration is in-
duced by the coculture is suggested by the absence of deterioration in control beads, which remained intact (data not
shown).
As early as two days, and throughout the remain incubation
period, coimmobilized C. vulgaris and P. myrsinacearum
were dying off as indicated by the presence of “ghost” cells
(empty cell envelopes) within the cavities (Figs. 1c, 2e). After ten days of incubation, many ghost cells of both microorganisms were observed (Fig. 2f, g). When immobilized alone
or with P. myrsinacearum, the senescent microalga increased
the number of grana, accumulated starch granules inside the
chloroplast, produced lipid granules (in the cytoplasm), and
increased the electron density of its nucleus (Figs. 1b, d; 2a,
d; 4 stage III). These ultrastructural events support an earlier
study in which P. myrsinacearum was unable to enhance the
growth of the microalga (Gonzalez-Bashan et al. 2000).
Most of the senescence events did not occur when the
microalga was coimmobilized with A. brasilense. This
PGPB may alter the metabolism of the microalga and delay
its senescence possibly through the production of plant
© 2001 NRC Canada
6
Fig. 4. Schematic model explaining the developing interactions between the wastewater-treatment microalga C. vulgaris and either its associative bacterium P. myrsinacearum or
the plant growth-promoting bacterium A. brasilense in alginate beads immersed in the mineral salts of Residual Water Medium as viewed ultrastructurally. a- interaction with
P. myrsinacearum, and b-interaction with A. brasilense. Each section of this model stands separately as the coculture (microalgae – bacteria) was never done with both bacteria.
Electron micrograph evidence for deterioration of the bead periphery was published before (Gonzalez-Bashan 2000).
Can. J. Microbiol. Vol. 47, 2001
© 2001 NRC Canada
Lebsky et al.
hormones, a well-established characteristic of A. brasilense
(Costacorta and Vandeleyden 1995; Patten and Glick 1996).
Azospirillum brasilense was previously shown to maintain
inoculated cardon cactus seedlings in the juvenile phase for
a longer-than-normal time (Puente and Bashan 1993), to
prolong the growth period of the seaweed Salicornia
bigelovii (Bashan et al. 2000), and to increase the population
of C. vulgaris (Gonzalez and Bashan 2000).
An electron-dense material of unidentified nature, formed
on the inner surface of the bead cavities concomitant with an
increase in the electron density of the cells in all
coimmobilized systems (Figs. 1b, c; 2a, d, e; 3c). Despite
the senescence of C. vulgaris cells when coimmobilized
with P. myrsinacearum, dividing microalgal and bacterial
cells appeared frequently in the culture (Fig. 2c, e; large arrows indicate dividing cells).
The ultrastructural evidence presented in this study allows
us to propose the following hypothetical model for the interaction between C. vulgaris and plant associative bacteria in
alginate beads (Fig. 4). The coimmobilization process and
the initial multiplication periods are similar for both bacterial species as described above (stages I and II). Longer incubations differentiate between the response of C. vulgaris
toward the two bacterial partners; senescence in the presence
of its associative bacterium and multiplication in the presence of the plant growth-promoting bacteria (stage III, a, b).
In summary, this paper provides ultrastructural evidence for
possible commensal interactions between the microalga
C. vulgaris and two plant associative bacteria when
coimmobilized in alginate beads, a preparation proposed for
advanced wastewater treatment.
Acknowledgements
Y. Bashan participated in this study in memory of the late
Mr. Avner Bashan from Israel. We thank Dr. Elena Morzhina,
Mr. Vladimir Semjonov, and Mr. Sergej Stephanov from the
Institute of Agricultural Microbiology, St. PetersburgPushkin, Russia for technical help in electron microscopic
observations, Mr. Angel Carrillo for artwork, Dr. Ellis Glazier for editing the English-language text, and Mrs. Cheryl
Patten for the English style. This study was supported by
Consejo Nacional de Ciencia y Tecnología (CONACyT),
Mexico, contracts #26262-B and #28362-B, by Instituto
Colombiano para el Desarrollo de la Ciencia y la
Tecnología, Francisco José de Caldas (COLCIENCIAS), Colombia, and by the Bashan Foundation.
References
Assmus, B., Hutzler, P., Kirchhof, G., Amann, R., Lawrence, J.R.,
and Hartmann A. 1995. In situ localization of Azospirillum brasilense in the rhizosphere of wheat with fluorescently labeled,
rRNA-targeted oligonucleotide probes and scanning confocal laser microscopy. Appl. Environ. Microbiol. 61: 1013–1019.
Baca, B.E., Soto-Urzua, L., Xochihua-Corona, Y.G., and CuervoGarcia, A. 1994. Characterization of two aromatic amino acid
aminotransferases and production of indoleacetic acid in
Azospirillum strains. Soil Biol. Biochem. 26: 57–63.
Bashan, Y. 1986. Alginate beads as synthetic inoculant carriers for
the slow release of bacteria that affect plant growth. Appl. Environ. Microbiol. 51: 1089–1098.
7
Bashan, Y. 1998. Inoculants of plant growth-promoting bacteria for
use in agriculture. Biotechnol. Adv. 16: 729–770.
Bashan, Y., and Gonzalez, L.E. 1999. Long-term survival of the
plant-growth-promoting bacteria Azospirillum brasilense and
Pseudomonas fluorescens in dry alginate inoculant. Appl.
Microbiol. Biotechnol. 51: 262–266.
Bashan, Y., and Holguin, G. 1994. Root-to-root travel of the beneficial bacterium Azospirillum brasilense. Appl. Environ.
Microbiol. 60: 2120–2131.
Bashan, Y., and Holguin, G. 1997a. Azospirillum-plant relationships: Environmental and physiological advances (1990–1996).
Can. J. Microbiol. 43: 103–121.
Bashan, Y., and Holguin, G. 1997b. Short and medium term avenues for Azospirillum inoculation. In Plant Growth-Promoting
Rhizobacteria- present status and future prospects. Edited by A.
Ogoshi, K. Kobayashi, Y. Homma, F. Kodama, N. Kondo, and S.
Akino. Faculty of Agriculture, Hokkaido University, Sapporo,
Japan. pp. 130–149.
Bashan, Y., and Holguin, G. 1998. Proposal for the division of
Plant Growth-Promoting Rhizobacteria into two classifications:
Biocontrol-PGPB (Plant Growth-Promoting Bacteria) and
PGPB. Soil Biol. Biochem. 30: 1225–1228.
Bashan, Y., and Levanony, H. 1989. Factors affecting adsorption of
Azospirillum brasilense Cd to root hairs as compared with root
surface of wheat. Can. J. Microbiol. 35: 936–944.
Bashan, Y., Levanony, H., and Klein, E. 1986. Evidence for a weak
active external adsorption of Azospirillum brasilense Cd to
wheat roots. J. Gen. Microbiol. 132: 3069–3073.
Bashan, Y., Levanony, H., and Ziv-Vecht, O. 1987. The fate of
field-inoculated Azospirillum brasilense Cd in wheat
rhizosphere during the growing season. Can. J. Microbiol. 33:
1074–1079.
Bashan, Y., Levanony, H., and Whitmoyer, R.E. 1991. Root surface
colonization of non cereal crop plants by pleomorphic
Azospirillum brasilense Cd. J. Gen. Microbiol. 137: 187–196.
Bashan, Y., Holguin, G., and Lifshitz, R. 1993. Isolation and characterization of plant growth-promoting rhizobacteria. In
Methods in plant molecular biology and biotechnology. Edited
by B.R. Glick and J.E. Thompson. CRC Press, Boca Raton,
Florida. pp. 331–345.
Bashan, Y., Moreno, M., and Troyo, E. 2000. Growth promotion of
the seawater-irrigated oilseed halophyte Salicornia bigelovii inoculated with mangrove rhizosphere bacteria and Azospirillum
spp. Biol. Fertil. Soils, 31: (in press).
Berg, R.H., Vasil, V., and Vasil, I.K. 1979. The biology of
Azospirillum-sugarcane association. II. Ultrastruct. Protopl. 101:
143–163.
Bettmann, H., and Rehm, H. 1984. Degradation of phenol by polymer
entrapped microorganisms. Appl. Microbiol. Biotechnol. 20:
285–290.
Cassidy, M.B., Lee, H., and Trevors, J.T. 1996. Environmental applications of immobilized microbial cells: A review. J. Ind.
Microbiol. 16: 79–101.
Chevalier, P., and De la Noüe, J. 1988. Behavior of algae and bacteria co-immobilized in carrageenan, in a fluid bed. Enzymol.
Microb. Technol. 10: 19–23.
Costacurta, A., and Vanderleyden, J. 1995. Synthesis of phytohormones by plant-associated bacteria. Crit. Rev. Microbiol. 21:
1–18.
De la Noüe, J., and Pauw, N. 1988. The potential of microalgal
biotechnology: A review of production and uses of microalgae.
Biotechnol. Adv. 6: 725–770.
Dyrset, N., Lystad, K.Q., and Levine, D.W. 1994. Development of
a fermentation process for production of an alginate G-lyase
© 2001 NRC Canada
8
from Klebsiella pneumoniae. Appl. Microbiol. Biotechnol. 41:
523–530.
Fages, J. 1992. An industrial view of Azospirillum inoculants: Formulation and application technology. Symbiosis, 13: 15–26.
Gonzalez, L.E., and Bashan, Y. 2000. Increased growth of the
microalgae Chlorella vulgaris when coimmobilized and
cocultured in alginate beads with the plant growth-promoting
bacterium Azospirillum brasilense. Appl. Environ. Microbiol.
66: 1527–1531.
Gonzalez, L.E., Cañizares, R.O., and Baena, S. 1997. Efficiency of
ammonia and phophorus removal from Colombian
agroindustrial wastewater by the microalgae Chorella vulgaris
and Scenedesmus dimorphus. Biores. Technol. 60: 259–262.
Gonzalez-Bashan, L.E., Lebsky, V.K., Hernandez, J.P., Bustillos,
J.J., and Bashan, Y. 2000. Changes in the metabolism of the
microalga Chlorella vulgaris when coimmobilized in alginate
with the nitrogen-fixing Phyllobacterium myrsinacearum. Can.
J. Microbiol. 46: 653–659.
Kayano, H., Matsunaga, T., Karube, I., and Suzuki, S. 1981. Hydrogen evolution by co-immobilized Chlorella vulgaris and
Clostridium butyricum cells. Biochem. Biophys. Acta, 638: 80–85.
Khang, Y.H., Shankar, H., and Senatore, F. 1988. Enhanced betalactam antibiotic production by coimmobilization of fungus and
algae. Biotechnol. Lett. 10: 867–872.
Knösel, D.H. 1962. Prüfung von bakterien auf Fähigkeit zur
Sternbildung. Zentralbl. Bakteriol. Parasitenk. Infecktionskr.
Hyg. Abt. 2, 116: 79–100.
Knösel, D.H. 1984. Genus Phylobacterium. In Bergies manual of
systematic bacteriology. Edited by N.R. Kreig, and J.G. Holt.
Vol. 1, The Williams and Wilkins Co. Baltimore. pp. 254–256.
Lambert, B., Joos, H., Dierickx, S., Vantomme, R., Swings, J.,
Kersters, K., and Van Montagu, M. 1990. Identification and
plant interaction of a Phyllobacterium sp., a predominant
rhizobacterium of young sugar beet plants. Appl. Environ.
Microbiol. 56: 1093–1102.
Lau, P.S., Tam, N.F.Y., and Wong, Y.S. 1997. Wastewater nutrients
(N and P) removal by carrageenan and alginate immobilized
Chlorella vulgaris. Environ. Technol. 18: 945–951.
Leenen, E.J.T.M., Dos Santos, V.A.P., Grolle K.C.F., Tramper, J.,
and Wijffels, R.H. 1996. Characteristics of and selection criteria
for support materials for cell immobilization in wastewater treatment. Water Res. 30: 2985–2996.
Levanony, H., Bashan, Y., Romano, B., and Klein, E. 1989.
Ultrastructural localization and identification of Azospirillum
brasilense Cd on and within wheat root by immuno-gold labeling. Plant Soil, 117: 207–218.
Can. J. Microbiol. Vol. 47, 2001
Mchugh, D.J. 1987. Production, properties and uses of alginates. In
Production and utilization of products from seaweeds. Edited by
D.J. Mchugh. Food and Agriculture Organization of the UN,
Rome. pp. 58–115.
Oh-Hama, T., and Miyachi, S. 1992. Chlorella. In Micro-algal biotechnology. Edited by M.A. Borowitzka and L.J. Borowitzka.
Cambridge University Press. Cambridge, U.K. pp. 3–26.
Okon, Y., and Labandera-Gonzalez, C.A. 1994. Agronomic applications of Azospirillum: An evaluation of 20 years worldwide
field inoculation. Soil Biol. Biochem. 26: 1591–1601.
Omar, S.H. 1993. Oxygen diffusion through gels employed for immobilization. 2. In the presence of microorganisms. Appl.
Microbiol. Biotechnol. 40: 173–181.
O’Reilly, A.M., and Scott, J.A. 1995. Defined coimmobilization of
mixed microorganism cultures. Enzymol. Microb. Technol. 17:
636–646.
Patten, C.L., and Glick, B.R. 1996. Bacterial biosynthesis of
indole-3-acetic acid. Can. J. Microbiol. 42: 207–220.
Puente, M.E., and Bashan, Y. 1993. Effect of inoculation with
Azospirillum brasilense strains on the germination and seedlings
growth of the giant columnar Cardon cactus (Pachycereus
pringlei). Symbiosis, 15: 49–60.
Puente, M.E., Holguin, G., Glick, B.R., and Bashan, Y. 1999. Root
surface colonization of black mangrove seedlings by
Azospirillum halofraeferens and Azospirillum brasilense in seawater. FEMS Microbiol. Ecol. 29: 283–292.
Schank, S.C., Smith, R.L., Weiser, G.C., Zuberer, D.A., Bouton,
J.H., Quesenberry, K.H., Tyler, M.E., Milam, J.R., and Littell,
R.C. 1979. Fluorescent antibody technique to identify
Azospirillum brasilense associated with roots of grasses. Soil
Biol. Biochem. 11: 287–295.
Tam, N.F.Y., Lau, P.S., and Wong, Y.S. 1994. Wastewater inorganic
N and P removal by immobilized Chlorella vulgaris. Water Sci.
Technol. 30: 369–374.
Tanaka, A., and Nakajima, H. 1990. Applications of immobilized
growing cells. Adv. Biochem. Eng. Biotechnol. 42: 97–131.
Uchida, M., and Nakayama, A. 1993. Isolation of Laminaria-frond
decomposing bacteria from Japanese coastal waters. Bull. Jap.
Soc. Sci. Fish, 59: 1865–1871.
Vande Broek, A., Michiels, J., Van Gool, A., and Vanderleyden, J.
1993. Spatial-temporal colonization pattern of Azospirillum
brasilense on the wheat root surface and expression of the bacterial nifH gene during association. Mol. Plant-Microbe Interact.
6: 592–600.
Wikstrom, P., Szwajcer, E., Brodelius, P., Nilsson, K., and Mosbach,
K. 1982. Formation of keto acids from amino acids using immobilized bacteria and algae. Biotechnol. Lett. 4: 153–158.
© 2001 NRC Canada