EL-SEVIER
FEMS Microbiology Ecology 16 (1995) 185-192
Phytoplankton
cell size and the development of
microenvironments
Laurie L. Richardson
a,*, Keith D. Stolzenbach
b
a Department of Biological Sciences, and Drinking Water Research Center, Florida International Uniuersity, Miami, FL 33199, USA
b Civil and Environmental
Engineering,
University of California, Los Angeles, CA 90024, USA
Received 21 July 1994; revised 11 October 1994; accepted 24 October 1994
Abstract
The effect of cell size on the development of extracellular microenvironments
of pH produced by individual photosynthesizing phytoplankton cells was investigated. The presence of pH microenvironments
was determined by detection of a
chemical reaction known to occur in microenvironments
of high pH produced by algal photosynthesis,
specifically the
extracellular oxidation of M&I) to MnO,. The dye leukoberbelin blue was used to detect the reaction. It was experimentally
determined that individual algal cells larger than 20 pm (length and/or width) produced microenvironments, while smaller
cells did not unless present as cell aggregates larger than 20 pm. A mathematical model is presented and discussed.
Keywords:
Microenvironments;
Phytoplankton;
Manganese
1. Introduction
Several reports have documented the existence of
microenvironments
of 0, and/or pH associated with
marine snow [ll and aggregates of phytoplankton
[2,3] in pelagic aquatic environments.
In each case
the microenvironments
were the result of biological
activity, specifically 0, consumption during respiration [1,2], and elevation of pH (due to CO, depletion) and 0, evolution during photosynthesis [3].
A question which arises from these results is that
of the effect of cell size. Are chemical microenvironments limited to aggregates of cells (bacterial or
* Corresponding author. Fax: (1) (305) 348-3894. E-mail:
[email protected].
0168.6496/95/$09.50
algal) on the order of millimeters (the size range of
phytoplankton
aggregates and marine snow particles
investigated),
or can individual cells generate microenvironments?
Since it was shown that the
microenvironments
were the result of cellular
metabolism
[l-3],
theoretically,
photosynthesis
or
respiration by an individual cell could generate a
microenvironment.
In the cases cited above, chemical microenvironments
were measured directly by
means of pH and 0, micro- or mini-electrodes,
commonly used in microbial ecology [4]. Detection
of microenvironments
produced by individual cells,
however, would be difficult or impossible using the
microelectrode
technique, because the size of the
electrode tip is comparable to, or larger than, the size
of individual cells. The smallest tips are on the order
of 3 pm in diameter, larger than many bacteria and
in the size range of small phytoplankton
cells. In
0 1995 Federation of European Microbiological Societies. AI1 rights reserved
SSDI 0168-6496(94)00082-4
186
L.L. Richardson, K.D. Stolzenbach / FEM.7 Microbiology Ecology 16 1199s) 185-192
addition, minimum spatial resolution obtainable using microelectrodes is on the order of 50 pm due to
diffusive limitations. Therefore, investigation of development of microenvironments
by individual cells
requires use of alternative methods.
One method is the use of dyes which specifically
detect a compound associated with the microenvironment. This approach was used by Paerl and Prufert
[5], where they used tetrazolium salts to detect areas
of low oxidation/reduction
potential associated with
bundles of filamentous cyanobacteria.
The dye approach was corroborated by direct measurement
of
anoxic zones using microelectrodes [5].
Here we report experimental
use of a dye to
detect extracellular pH microenvironments
at a much
smaller spatial resolution than can currently be resolved using microelectrodes.
Previous research by
Richardson et al. [3] demonstrated that aggregates of
phytoplankton
can oxidize Mn(I1) (the most reduced
form of manganese) to Mn(II1) and M&V), which
form insoluble oxides generally designated as MnO, .
The Mn oxidation reaction is specifically due to high
pH produced in microenvironments
surrounding the
aggregates [3], the result of photosynthetic
removal
of CO, 161. While 0, (which is required for the
reaction) concentration
also increases during algal
photosynthesis, favoring manganese oxidation, it was
shown [3] that the reaction is primarily due to generation of high pH. This was demonstrated by showing
that inhibition of manganese oxidation occurred when
pH was adjusted and maintained at or below 8.0,
even though 0, evolution by photosynthesis continued [3]. These results are consistent with Stumm and
Morgan’s rate equations for Mn(II) oxidation [6],
where M&I) oxidation rate is second order with
respect to pH, but only first order with respect to
[O,]. According to Stumm and Morgan [6], Mn(II)
oxidation rate rapidly accelarates at pH values above
9.0, and at pH values above 10.0 (with 1 atm of O,),
manganese oxidation becomes autocatalytic.
Direct
measurement (using a minielectrode) of pH microenvironments produced by aggregates of phytoplankton
which were oxidizing M&I) revealed pH values up
to 10.75 [3]. The oxidation of M&I) was prevented
by adjusting pH to 8.0 or below, by incubating in
darkness, or by addition of the poison DCMU (3(3,4)-dichlorophenyl-(l,l)-dimethylurea)
which
specifically
inhibits photosynthesis
[3]. The man-
ganese oxidation mechanism was later corroborated
by Lubbers et al. [7] in the North Sea, and has been
adopted to explain manganese deposition in the microfossil record [8].
To investigate the effect of individual algal cell
size on generation of microenvironments,
we used
the dye leukoberbelin blue, an indicator which turns
from colorless to blue when in contact with manganese of oxidation state 3 or above [9]. The effect
of algal cell size on generation of high pH microenvironments was studied by detection of the production of manganese oxides, which occurs specifically
within the high pH microenvironments.
2. Materials and methods
Algal species used in all experiments were isolated into axenic culture, and maintained in laboratory stock cultures in liquid medium (medium ‘D’
[lo] for Cyanophyta and Chlorophyta, and Chu #lO
for Chrysophyta).
Cultures were routinely checked
for contamination
by heterotrophic bacteria by plating onto agar plates which contained yeast extract.
All cultures were obtained from Oneida Lake, New
York, a eutrophic freshwater lake. Algae investigated
in the laboratory were Ankistrodesmus sp., Microcystis sp., Chlorella sp., Nitzschia sp., Anabaena
sp., Scenedesmus sp., Vaucheria sp., and an unidentified unicellular
Chlorophyte
(Order Chlorococtales). Cell dimensions
were measured using an
ocular micrometer inserted into the eyepiece of a
compound microscope. The minimum cell dimension
(i.e. cell width or length) ranged from 3 to 50 pm.
Manganese oxidation by individual algal cells was
investigated in algal mineral medium ‘D’. Ten ml of
media, with 25 FM Mn(II) added from a stock
solution of MnCl,, were placed in a sterile petri dish
which contained a sterile microscope slide. (Controls
were incubated without added manganese, but contained 2.7 PM Mn as a source of this required trace
nutrient.) A suspension of the test alga, from axenic
cultures maintained
in the laboratory, was gently
pipetted over the microscope slide and allowed to
settle onto the slide. Slides were then inspected using
a dissecting microscope (viewing through the petri
dish cover to maintain sterile conditions) to ensure
that individual cells were not touching each other.
L.L. Richardson, K.D. Srolzenbach/FEMS
The cultures were incubated under low light (85
pEins/m*/s)
supplied by a cool-white fluorescent
bulb, for 4 days, under non-turbulent
conditions (i.e.
the petri dishes remained stationary). After this period, the medium was carefully pipetted from the
petri dish, allowing algal cells to remain on the slide
surface. A solution of leukoberbelin
blue (0.04%
w/v in 0.25% acetic acid, prepared as described by
Krumbein and Altmann [9]), was then pipetted carefully onto the microscope slide. When oxidized manganese was present, revealing that manganese oxidation had occurred, the solution turned blue. Slides
were also viewed using a compound microscope. A
blue reaction was scored as positive. No color change
was scored as negative.
In addition to investigating
individual
cells in
dilute suspensions
as detailed above, experiments
were conducted using aggregates of cells. These
were carried out in the same manner with the exception that inoculation
of the slide was done using
much higher concentrations
of culture material, manipulated during pipetting to allow deposition of
aggregates of cells (confirmed by microscopy)
as
opposed to individual cells. During scoring of these
test algae, the size of the aggregate was measured
using an ocular micrometer.
Besides the controls incubated without the addition of manganese,
parallel incubations
were run
with the medium maintained at pH 8.0 using 10 mM
HEPES buffer, or with the addition of 5 PM DCMU.
3. Results and discussion
Results of these experiments are shown in Table
1. Cells with at least one dimension (width or length)
of 20 pm or larger consistently oxidized Mn(I1) to
MnO, (i.e. scored positive in the leukoberbelin blue
test), while smaller cells did not. Of the smaller cells
investigated, one species of Nitzschia had lengths up
to 18 pm, yet was never observed to oxidize manganese. When cells smaller than 20 pm were
clumped into aggregates, the aggregates oxidized
manganese (Table 1). Both the addition of 5 PM
DCMU and adjusting the pH to 8.0 (using HEPES
buffer) prevented manganese oxidation in all ‘positive’ test cultures, which corroborated
that manganese oxidation was driven by photosynthetically
Microbiology
Ecology 16 (1995) 185-192
187
elevated pH as shown previously [3]. No oxidation
was detected in experiments with lo-fold lower concentration
of M&I)
(2.7 PM) during the 4-day
incubation period.
Microscopic analysis after addition of leukoberbelin blue revealed that, in addition to the color reaction, ‘positive’ individual cells exhibited a thin blue
coating on the cell surface. The cells looked normal,
but were blue. Aggregates, however, exhibited distinct extracellular accumulations of particulate material, which turned dark blue when leukoberbelin blue
was added. The accumulation of extracellular material did not occur when pH was adjusted to 8.0, or
when DCMU was added. We interpret this as evidence that the particulate matter is MnO,. These
results are consistent with earlier findings of extracellular manganese oxidation, in which manganese
oxidation (conversion of manganese from soluble to
particulate) was also documented using atomic absorption spectroscopy [3]. An example of the extracellular production of MnO, is shown at a high
magnification in Fig. 1.
The experiments described above were conducted
under motionless, i.e. non-stirred or agitated, conditions. While natural aquatic ecosystems may at times
be non-turbulent,
this is not the norm. Turbulence
has been shown to be important in the development
and dynamics of microenvironments
of oxygen and
pH [ll]. No attempt was made to repeat these experiments and investigate the effect of water movement,
and no experiments were incubated in situ. However,
freshly collected phytoplankton samples from Oneida
Lake, New York, were tested with leukoberbelin
blue to determine if associated oxidized manganese
was present, and if the size ranges of samples with
positive reactions were consistent with the laboratory
results. In addition to this field study, a literature
survey of published reports of manganese oxides
associated with algae was carried out. In each report,
cell sizes were noted if reported, or determined from
the literature, again with the goal of determining if
algal size was in agreement with our laboratory
results.
Table 2 presents data retrieved from the literature,
and results of the field study carried out at Oneida
Lake (in collaboration
with E. Mills), of oxidized
manganese
associated with phytoplankton.
In all
cases these data are consistent with our laboratory
188
L.L. Richardson, K.D. Stolzenbach/FEMS
Microbiology
Ecology 16 (1995) 185-192
Fig. 1. Scanning electron micrograph of a culture of Nitzschia sp. with extracellular, particulate
when aggregates of cells were present, as is shown. Frustules measured 3 by 18 pm. ( X 4000).
Table 1
Generation
of size a
of a pH microenvironment
Cell investigated
(detected by oxidation
Size ( pm)
of manganese)
sp.
Microcystis sp.
Chforella sp.
Nitzschia sp.
Anabaena
sp.
Scenedesmus
sp.
Unicellular green
( Vaucheria sp.) ’
a Controls are not
b Aggregates were
20 pm and larger
’ This genus is an
width
length
diam.
diam.
width
length
width
length
width
length
diam.
width
length
= 3
= 12
= 4
= 9
= 3
to 18
= 3
= 20 +
= 10
= 20
= 20
= 50
= mm’s
phytoplankton
MnO, formed
Cell
Ankistrodesmus
by individual
manganese
oxides. Oxidation
cells and aggregates
Taxonomic
Aggregate
as a function
group
’
+
Chlorophyta
+
+
+
Cyanophyta
Chlorophyta
Chrysophyta
+
Cyanophyta
+
Chlorophyta
+
+
Chlorophyta
Chrysophyta
included here, and are discussed in the text. All controls were negative.
of variable size depending on the number of cells in the aggregate and size of cells. Aggregates were measured,
were positive.
attached, filamentous alga and not a phytoplankter, but was included in the study due to its large size.
and those
L.L. Richardson, K.D. Stol.zenbach/FEMS
findings. Of particular interest are the reports by
Hunt and Smith [12] and Lubbers et al. [7]. Hunt and
Smith reported conversion of dissolved, soluble manganese, Mn(II1, to particulate (oxidized) manganese
in conjunction
with a bloom of ‘large’ diatoms in
MERL microcosms. The species of diatoms present
in the report were not recorded, and for Table 2
general size ranges are given which were taken from
the literature. The retrieved size ranges are in agreement with our laboratory data, with all species having a length or width of at least 30 pm.
The study by Hunt and Smith [12] consists of a
report of a large-scale manganese precipitation event
in MERL mesocosms. During their study, the presence of the ‘large’ diatoms resulted in mass conversion of dissolved to particulate manganese with accompanying net flux of manganese to the sediments.
When ‘smaller’ phytoplankton
were present (microflagellates), there was a net outflow of soluble (re-
Table 2
Survey of oxidized manganese
43
acritarchs
Phaeocystis
sp. b
associated
with natural phytoplankton
189
duced) manganese from the mesocosm system. While
the mechanism of the event was unexplained,
the
data are compatible with the microenvironment
oxidation mechanism.
Also included in Table 2 is a study by Lubbers et
al. [7] which documented manganese oxidation by
colonies of Phaeocystis sp. in the North Sea (a paper
which corroborated oxidation in high pH microenvironments). This paper did not report the size of the
cells, however individual
Phaeocystis cells (which
did not oxidize manganese)
were separated from
Phaeocystis colonies (which did oxidize manganese)
by 20 pm plankton gauze [7], again consistent with
our findings.
Another study [8] noted in Table 2 documented
the association
of manganese
oxides with fossil
acritarchs, which are microfossils of planktonic plant
protists believed to be related to Modem phytoplankton. This report interpreted the presence of MnO, to
blooms and fossil microalgae
Size ( pm)
MnO, Associated
t81
30
< 20 (cells)
> 20 (aggregates)
40-130
40-600
4 (cells)
> 20 (aggregates)
width = 3
length = 20
width = lo-30
length = variable
described in paper as ‘large’
10 by 200-450
4-8 by 60-150
width = 50
length = variable
(see footnote)
+
+
+
+
_
+
+
+
+
+
+
+
+
+
+
-
[71
(Mills and Richardson,
(Mills and Richardson,
Anabaena sp.
[31
Chaetoceros
1121c
[31
Chaetoceros sp.
Nitzschia longissima
Baciliaria paradoxa
Lithodesmium undulatum
I121
1121
1121
1121
Microflagellates
1121
d
Ecology 16 (1995) 185-192
Reference
Asterionella sp.
Fragilaria sp.
Microcystis sp.
curvisetus
Microbiology
unpublished)
unpublished)
a
a In this category, the conversion of dissolved to particulate manganese in MERL mesocosms is assumed to be due to oxidation.
b The dimensions of the Phaeocystis sp. reported in this paper were not given, however, within the study, colonies were separated from
individual cells by 20 pm plankton gauze - therefore, cells were less than 20 pm.
’ The dimensions of these species were not given in this paper but were taken from the literature as follows. C. curuisetus, L. undulatum,
from Hustedt, F. (1930) Die Kiesalalgen, Otto Koeltz Science Publ., Koenigstein, West Germany; N. longissima, from Hendey, N.I. (1964)
An introductory Account of the Smaller Algae of British Coastal Waters, Part V. Bacillariophyceae.
Her Majestey’s Stationery Office,
London; B. paradoxa, from Werner, D. (1977) The Biology of Diatoms, Univ. California Press.
d The dimensions of the microflagellates
were not given, nor were those of taxon. However, in general microflagellates
are considered to be
only a few microns wide.
190
L.L. Richardson, K.D. Stolzenbach/ FEMS Microbiology Ecology 16 (1995) 185-I 92
be due to extracellular manganese oxidation in high
pH microenvironments,
again corroborating our earlier work [3]. The size of the acritarchs was reported
to be 30 pm in diameter [S].
The data in both Tables 1 and 2 show that manganese oxidation was consistently
associated with
algal cells, colonies, or aggregates which were larger
than 20 pm.
Our experimental,
and literature survey, findings
of the correlation of manganese oxide formation with
cell size is consistent with the physics of CO, transport (to support photosynthetic CO, fixation) which
occurs at the cell surface. It is well known that a
solution of CO, in water will equilibrate, via several
dissociation
reactions, to a mix of the inorganic
carbon species H,CO,,
HCO;,
CO:and CO,,
which buffers the pH of the system [6,13]. It is also
well known that removal of CO, in aquatic systems
increases pH [6,13]. The relationships
among cell
size, carbonate uptake at the cell surface, elevation
of surface pH, (and potential manganese precipitation) could theoretically be modeled. Such a mathematical analysis would have to consider which of
several carbonate species the cell was taking up, the
kinetics of reactions between different carbonate
species in the microzone around the ceil, and diffusion of acid/base
species towards and away from
the surface of the cell [14].
In a motionless fluid environment, uptake of carbon by a nearly spherical photosynthesizing
cell will
result in a localized decrease in the inorganic carbon
concentration at the cell surface given by:
’ 67. If these two dependencies of cell properties on cell size are assumed to remain constant,
the decrease in total carbon AC, increases approximately linearly with cell size R.
The uptake of CO, is related to photosynthetic
rate, which in turn is related to cell biomass. The
generation of a pH microenvironment
at the cell
surface, the result of cellular uptake, is also controlled by diffusion of acidic species to the surface of
the cell, and of basic species away from the cell
(with both acidic and basic species present within the
carbonate equilibria [6]). The diffusion/size
relationship is expressed as:
diffusion
= 47rRDAC,
where AC is the change in concentration of diffusing species between the cell surface and the surrounding solution. Diffusion is proportional to R.
Thus as the diameter of the cell increases, with
uptake increasing proportional to R2, diffusion of
acidic and basic carbon species will, at some point,
not be able to equilibrate with uptake of CO, and a
microzone of elevated pH will be established at the
cells surface.
Extremely rapid manganese oxide precipitation,
which occurs whenever AC, is large enough to
elevate the pH above 9.0 [6], will thus be restricted
to larger cells. This theoretical result would not be
changed by consideration of the effect of fluid motion (sinking or fluid turbulence) on mass transfer to
the cell, although the theoretical dependency of AC,
on R would be somewhat weaker [19].
A predictive mathematical model would necessitate, in addition to dissolved inorganic carbonate
chemistry and diffusion of chemical species, incorporation of variation in cell metabolism, including
rates of photosynthesis
and respiration,
effect of
changing environmental parameters such as light and
L.L. Richardson, K.D. Stolzenbach/FEMS
temperature
on metabolic rate, uptake/release
of
other ions and nutrients (which could also effect
extracellular pH), as well as the potential role of cell
surface chemistry.
The results presented here, supported by mathematical considerations,
show that individual
algal
cells can and do generate chemical microenvironments which can be biogeochemically
significant.
Acknowledgements
We would like to thank John Raven, Francois
Morel, and Janet Hering for helpful discussions, and
John Makemson for comments on the manuscript.
Two anonymous
reviewers greatly improved the
manuscript and their input is very much appreciated.
We would also like to thank Ed Mills for collaborative field research at the Cornell Biological Field
Station. This research was supported by a National
Research Council Research Associateship,
and by
the National Atmospheric and Space Administration
(grant NAGW1047).
References
[I] Alldredge, A.L. and Cohen, Y. (1987) Can microscale chemical patches persist in the sea? Microelectrode
study of
marine snow, fecal pellets. Science 235, 689-691.
[Z] Paerl, H.W. and Bebout, B.M. (1988) Direct measurement of
O,-depleted
microzones in marine Oscillatoria: Relation to
N, fixation. Science 241, 442-445.
[3] Richardson,
L.L., Aguilar, C. and Nealson, K.H. (1988)
Manganese oxidation in pH and 0, microenvironments
produced by phytoplankton.
Limnol. Oceanogr. 33, 352-363.
141 Revsbech, N.P. and Jorgensen, B.B. (1986) Microelectrodes:
Their use in microbial ecology. Adv. Microb. Ecol. 9, 293352.
[s] Paerl, H.W. and Prufert, L.E. (1987) Oxygen-poor
microzones as potential sites of microbial N, fixation in nitrogen-
Microbiology
Ecology I6 (1995) 185-192
191
depleted aerobic marine waters. Appl. Env. Microbial. 53,
1078-1087.
[61 Stumm, W. and Morgan, J.J. (1981) Aquatic Chemistry,
Wiley, NY.
[71 Lubbers, G.W., Gieskes, W.W.C., de1 Castillo, P., Salomons,
W. and Bril, J. (1990) Manganese accumulation in the high
pH microenvironment
of Phaeocystis sp. (Haptophyceae)
colonies from the North Sea. Mar. Ecol. Prog. Ser. 59,
285-293.
@I Morad, S. and Vidal, G. (1989) Proterozoic Mn-oxide precipitation by planktonic plant protists (acritarchs). Geol. Mag.
126, 301-305.
191 Krumbein, W.E. and Altmann, H.J. (1973) A new method for
the detection and enumeration of manganese oxidizing and
reducing microorganisms.
Helgol. wiss. Meeresunters.
25,
347-356.
[lOI Castenholz, R.W. (1981) Isolation and cultivation of thermophilic cyanobacteria.
In: The Prokaryotes (Starr, M.P. et
al., Eds.), pp. 236-246, Springer.
and
[ill Carlton, R.G. and Wetzel, R.G. (1987) Distributions
fates of oxygen in periphyton communities. Can. J. Bot. 65,
1031-1037.
[121 Hunt, C.D. and Smith, D.L. (1980) Conversion of dissolved
manganese to particulate manganese during a diatom bloom:
Effects on the manganese cycle in the MERL microcosms.
In: Microcosms in Ecological Research, (Giesy, J.P., Ed.),
pp. 850-868. U.S. Tech. Information Center, U.S. Dept. of
Energy, Symp. Series 52 (CONF-781101).
[131 Wetzel, R.G. 1975. Limnology. W.B. Saunders Company,
Philadelphia, London, Toronto.
D.A. and Smetacek, V. (1993)
[141 Riebesell, U., Wolf-Gladrow,
Carbon dioxide limitation of marine phytoplankton
growth
rates. Nature 361, 249-251.
1151 Pasciak, W.J. and Gavis, J. (1974) Transport limitation of
nutrient uptake in phytoplankton,
Limnol. Oceanogr.
19,
881-888.
[I61 Morel, F.M.M. and Hering, J.G. (1993) Principles and Applications of Aquatic Chemistry. Wiley, NY.
[I71 Mullin, M.M., Sloan, P.R., and Eppley, R.W. (1966) Relationship between carbon content, cell volume, and area in
phytoplankton.
Limnol. Oceanogr. 11, 307-311.
k31 Nielsen, S.L. and Sand-Jensen, K. (1990) Allometric scaling
of maximal photosynthetic
growth rate to surface/volume
ratio. Limnol. Oceanogr. 35, 177-181.
[191 Jackson, G.A. (1989) Simulation of bacterial attraction and
adhesion to falling particles in an aquatic environment. Limnol. Oceanogr. 34, 514-530.