A novel approach to assess the respiration rate of microorganisms: case of the Chlorophycae Dunaliella tertiolecta (Butcher)

Gérald Grégori¹*, Michel Denis¹, Dominique Lefèvre¹ & Beatriz Beker²

¹ Laboratoire de Microbiologie, Géochimie et d'Ecologie Marines, Université de la Méditerranée, CNRS UMR 6117, 163 avenue de Luminy, Case 901, 13288 Marseille cedex 9, France;

² Centre d'Océanologie de Marseille, CNRS UMS 2196, Station Marine d'Endoume, rue de la Batterie des Lions, 13007 Marseille, France

*Present address : Purdue University Cytometry Laboratories, Hansen Life Sciences Research Building, B050, 201 S. University Street, West Lafayette, IN 47907-2064,USA

Corresponding author: gregori@flowcyt.cyto.purdue.edu

Introduction

The production of organic matter in the ocean is essentially the result of photosynthesis, a processus that has been intensively investigated for many years. The main fraction of this organic matter is transformed through the microbial trophic network and contributes to the energy transfer to higher trophic levels. The biological oxidation of organic matter proceeds through a variety of mechanisms that all converge towards the cell respiratory system. Under aerobic conditions that represent the most common situation throughout the water column, the last step of the electron transfer through the respiratory system consists in molecular oxygen reduction, perceived as oxygen uptake, coupled to metabolic production of carbon dioxide (CO₂) and synthesis of adenosine triphosphate (ATP), the cell energy fuel. The mineralisation of organic matter is mostly due to microorganisms, and in the aphotic layer, to heterotrophic bacteria. The resulting annual metabolic CO₂ production over the whole ocean was estimated to be around 22 Pg C y^-1 (Packard et al., 1988). Despite this large value of oceanic budget, the instant rate of metabolic CO₂ production is difficult to quantify by direct measurements and is usually estimated by determining the cellular respiration rate. This is achieved by indirect methods due to the low level of respiratory activity in oceanic waters, keeping the oxygen uptake out of reach of currently available direct techniques. Thus, bulk changes in dissolved oxygen concentrations (Winkler method; Aminot, 1983) or in dissolved inorganic carbon (coulometry; [13]) are assessed after long-term incubations (usually 24 hours) for sea-surface samples. The lower activity level in the aphotic layer prevents the extension of this approach below the euphotic layer. The enzymatic assessment of cellular respiratory activity as developed by Packard (1971) enabled addressing mineralisation down to the deep ocean. However, the need to convert the potential respiratory activity determined by this very sensitive method into in situ oxygen uptake rate by means of laboratory established conversion coefficients fueled lasting controversies.
In the present paper, we propose to combine flow cytometry and the use of fluorescent probes sensitive to membrane potential to face this lack of direct measurement methods for determining in situ microbial respiratory activity. The development of this new approach was based on (i) the existence of a linear relationship between the oxygen flux and the proton electrochemical potential difference (delta µH+) with respect to both sides of the inner mitochondrial membrane in yeast mitochondria (Rigoulet et al., 1987) and (ii) the availability of fluorescent probes sensitive to delta µH+. The thus determined oxygen uptake rates can be expressed in terms of metabolic CO₂ production by using conversion coefficients (Redfield et al., 1963; Takahashi et al., 1985).

Materials and methods

Phytoplankton culture
The unicellular organism chosen for the present work was the Chlorophyceae Dunaliella tertiolecta (Butcher), strain CS-175 from the CSIRO Marine Research Culture Collection (Hobart, Australia). It was grown under axenic conditions on f/2 medium [10] at 20 °C, a salinity of 33 and with an irradiance of 100 µE m^-2 s^-1 provided by fluorescent OSRAM lamps with a 12 h light-dark cycle.
Cells were always collected in the exponential growth phase and kept in the dark in order to address dark respiration and to prevent O₂ production would photosynthesis be activated.

Fluorescent probe
To monitor the cellular respiration by flow cytometry we used the fluorescent dye carbocyanine dihexyloxacarbocyanine iodide (DiOC₆(3), Molecular Probes, USA), in 100 µM working solutions prepared in DMSO (Sigma Chemical Co., USA).

Uncoupler
The respiration rate of D. tertiolecta was gradually enhanced by stepwise addition of carbonyl cyanide m-chloro phenyl hydrazone (CCCP, Sigma, USA), a potent uncoupler of mitochondrial electron transfer and proton translocation. The CCCP working solution (3 mM) was prepared in ethanol. We allowed for 10 min incubation time to compensate for small uncoupling efficiency variations between different CCCP batches.

High-resolution respirometry
D. tertiolecta respiration rates were recorded with the high-resolution oxygraph OROBOROS (Paar, Innsbruck, Austria), equipped with two measurement chambers (Gnaiger et al., 1983; 1995; 1998). The temperature in the chambers was set at 20 °C and controlled by a device based on the Pelletier effect. Cell suspensions were homogenized with a magnetic stirrer (600 rpm), and left to equilibrate with the atmosphere in the open chamber before signal calibration. Oxygen concentration was computed from the partial pressure measured by the electrode. During oxygen uptake measurements, each chamber (3 cm³ working volume) was closed by a piston to prevent oxygen exchange with the atmosphere. Data were recorded on a desktop computer every 0.4 s by using the Datlab Acquisition software (Paar, Innsbruck, Austria). Data analysis was run with the Datlab Analysis software (Paar, Innsbruck, Austria) to calculate oxygen consumption rates (µM.h^-1), and correct them from the electrode shift. The O₂-concentration-measurement uncertainty with the OROBOROS oxygraph was 0.036 µM O₂ against 0.500 to 1.000 µM O₂ for other commercial instruments.

Flow cytometry
Single cell analyses were run with a CYTORON ABSOLUTE (ORTHO Diagnostic Systems) flow cytometer, equipped with an air-cooled argon laser (excitation wavelength at 488 nm). Each cell was characterized by 5 optical parameters: two scatter parameters, namely forward angle scatter (related to the particle size) and right angle scatter (related to cell structure and shape), and three fluorescence parameters related to emissions in the red (> 620 nm), orange (565-592 nm) and green (515-530 nm) wavelength ranges. Data were collected and stored in list-mode on a personal computer with the IMMUNOCOUNT software (ORTHO Diagnostic Systems). Due to the control of the sample volume by a microsyringe, this software can directly provide cell concentrations (cells mm^-3). The Winlist software (Verity Software, USA) was used to determine statistical data.

Results

Titration of D. tertiolecta respiration
To cover a meaningful range of oxygen uptake rates, the respiration rate of D. tertiolecta was enhanced by stepwise addition of increasing amounts of CCCP, an uncoupler whose effect results in stimulating electron transfer and consequently, dioxygen uptake (Heytler, 1981, Rottenberg & Wu, 1998). The observed maximum enhancement (3.8 fold) corresponded to fully uncoupled mitochondria. Increasing values of O₂ uptake rate by D. tertiolecta were determined with the oxygraph after addition of increasing amounts of CCCP. As illustrated by Figure 1, we found a linear relationship between CCCP concentration and O₂ uptake rate by D. tertiolecta.

Figure 1. Relationship between the rate of oxygen uptake by D. tertiolecta and CCCP concentration. The straight line reveals the existence of a linear relationship between oxygen uptake rate and CCCP concentration: y = 2.45x + 17.18; R² = 0.99; n = 9.

Staining of D. tertiolecta
Control experiments run with DiOC₆(3) concentrations < 100 µM ensured that the dye had no apparent toxic effect during the time course of the observations. Indeed, oxygen uptake rates of stained cells (47.1 ± 5.8 µmol O₂ dm^-3 h^-1) were found not significantly different (10 measurements; p < 0.05) from that (46.1 ± 2.1 µmol O₂ dm^-3 h^-1) in absence of dye.
We defined an optimal dye concentration at 30 nM DiOC₆(3) and optimum staining was ensured by incubating D. tertiolecta cells 30 min in presence of DiOC₆(3) before flow cytometric analysis. Staining of D. tertiolecta was controlled by epifluorescence microscopy (not shown).

Monitoring of respiration
To titrate the fluorescence of DiOC₆(3) stained D. tertiolecta cells, subsamples were pretreated with increasing amounts of CCCP and incubated 30 min with DiOC₆(3) at 30 nM final concentration before flow cytometric analysis. Addition of CCCP decreased delta µH+ which resulted in the decrease of the green fluorescence signal of the stained cells as illustrated in Figure 2. We thus obtained a linear relationship between CCCP concentration and the cell mean green-fluorescence intensity (Figure 2). When mitochondria were fully uncoupled, the cell mean-green fluorescence intensity was similar to that of cells without staining (about 4 a.u., Figure 2).

Figure 2. Relationship between D. tertiolecta cell mean green fluorescence intensity and CCCP concentration. Note that the dye was supplemented after CCCP addition. The straight line expresses a linear relationship between the cell mean green fluorescence intensity (in arbitrary units, au) and CCCP concentration: y = -12.82x + 16.82; R² = 0.997; n = 4.

Conversion scale
From the two linear relationships obtained independently between CCCP concentration and D. tertiolecta oxygen uptake rate on the one hand and green fluorescence of stained D. tertiolecta on the other hand, we can conclude to the existence of a linear relationship between D. tertiolecta oxygen uptake rate and its green fluorescence when stained with DiOC₆(3). The ensuing conversion scale can be easily constructed from two specific values (derived from the fully coupled and fully uncoupled states respectively) of D. tertiolecta oxygen uptake rate and the associated green fluorescence of cells in the exponential growth phase, stained with DiOC₆(3). Both oxygen uptake rates (17 and 64 fmol O2 cell^-1 h^-1 for cells in fully coupled and fully uncoupled states respectively) can be considered as specific features of D. tertiolecta in exponential growth phase. As long as fluorescence values are expressed in arbitrary units, the conversion scale will only be valid for the instrument with which it was established. The linear conversion scale for our flow cytometer is presented in Figure 3. The green fluorescence values were corrected from the residual signal recorded with unstained cells.

Figure 3. Linear conversion scale between the cell O₂ respiration rate and the cell mean green fluorescence intensity (in arbitrary units, au) for D. tertiolecta stained with 30 nM DiOC₆(3). The linear scale is based on two specific respiration rates of D. tertiolecta in exponential growth phase, corresponding to fully coupled and fully uncoupled states respectively, and the associated fluorescence intensities. The resulting linear equation, y = -3.25x + 64.09, is therefore only valid for the instrument in use. To make this relationship instrument-independent would require a standardization of fluorescence measurements.

Validation
To check the efficiency of the linear conversion scale, we conducted an experiment in which the oxygen uptake rate of D. tertiolecta was measured directly with subsamples supplemented with CCCP (10 min) then with DiOC₆(3) (30 min). Once the oxygen uptake rate was measured, the cell suspension was removed from the oxygraph chamber to be analyzed by flow cytometry. The cell mean green fluorescence for each respiratory stage was corrected from the residual signal recorded with unstained cells and then converted to oxygen uptake rate by using the conversion scale shown in Figure 3. Figure 4 shows that calculated and measured oxygen uptake rates are not significantly different within the experimental uncertainties (R² = 0.984; 21 measures).

Figure 4. Comparison between measured and calculated mean cell respirations. The reported D. tertiolecta cell mean respiration rates were measured directly with the oxygraph and derived from the green fluorescence recorded by flow cytometry by using the linear scale conversion plotted in Figure 6. The straight line demonstrates a tight correlation between both data sets (y = 0.96x, R² = 0.98, n = 21).

Discussion

The established linear relationship between oxygen uptake rate and membrane potential (through CCCP tuning, Figure 1) extends to the whole cell a finding reported for isolated purified yeast mitochondria (Rigoulet et al., 1987). It enables a simple and direct conversion of fluorescence signals into oxygen uptake rates (Figure 3).
By demonstrating the feasibility to derive the respiration rate of D. tertiolecta from flow cytometric measurements, we opened the way to the extension of this approach to the other microorganisms. Indeed, this approach is based on the chemiosmotic theory of Mitchell (1961) which accounts for the bioenergetics of all cells, whether eukaryotes or prokaryotes. However, if there is no theoretical argument to preclude the extension of the present approach, experimental difficulties may be encountered in the way ahead. Whatever these difficulties, the present study opens the way to a new, elegant and efficient way to quantify cell respiration and subsequently, mineralisation by microorganisms, a major challenge in oceanography.

Acknowledgements
The authors wish to thank S. Brown and D. Marie for their encouragements and precious advices at the early stage of this work. G. G. was the recepient of a doctoral fellowship from the Council of the Provence-Alpes-Côte-d'Azur Region. Figures were reproduced from Gregori et al. (2002) with kind permission of Kluwer academic Publishers.

Cited references

Aminot A (1983) Dosage de l'oxygène dissous. In: CNEXO, (ed). Manuel des analyses chimiques en milieu marin, pp 75-92. Brest.

Gnaiger E, Forstner H (1983) Polarographic sensors, Aquatic and physiological applications. Gnaiger E, Forstner H (eds). Berlin/Heidelberg/New York: Springer-verlag.

Gnaiger E, Steinlechner-Maran R, Méndez G, Eberl T, Margreiter R (1995) Control of mitochondrial and cellular respiration by oxygen. J Bioenerg Biomemb 27(6): 583-596.

Gnaiger E, Kuznetsov AV, Lasnig B, Fuchs A, Reck M, Renner K, et al. (1998) High-resolution respirometry - optimum permeabilization of the cell membrane by digitonin. In: Larsson C, Pahlman TL, Gistatsson L, (eds), Bio thermokinetics in the post genomic era, pp 89-95. Göteborg: Chalmers Reproservice.

Grégori G, Denis M, Lefèvre D, Beker B (2002) A flow cytometric approach to assess phytoplankton respiration. Methods in Cell Science 24: 99-106.

Guillard RRL, Ryther JH (1962) Studies of marine planktonic diatoms. Part 1: Cyclotella nana Hustedt and Detonula confervacea (Cleve) Gran. Can J Microbiol 8: 229-239.

Heytler PG (1981) Uncouplers of oxidative phosphorylation. In: Erecinska M, Wilson DF (eds), Inhibitors of mitochondrial function: Pergamon Press pp. 199-210.

Johnson KM, Sieburth JM (1987) Coulometric total carbon dioxide analysis for marine studies: automation and calibration. Mar. Chem. 21: 117-133.

Mitchell P. (1961) Nature (Lond.) 191: 144-148.

Packard T (1971) The measurement of respiratory electron transport activity in marine phytoplankton. J Mar Res 29: 235-244.

Packard T, Denis M, Rodier M, Gardfield P (1988) Deep ocean metabolic CO2 production calculations from ETS activity. Deep-Sea Res. 35(3): 371-382.

Redfield AC, Ketchum BH, Richards FA (1963) The influence of organisms on the composition of sea-water. In: Hill MN (ed), The Sea, pp 26-77. New York: Wiley-Interscience.

Rigoulet M, Guérin B, Denis M (1987) Modification of flow-force relationships by external ATP in yeast mitochondria. Eur J Biochem 168: 275-279.

Rottenberg H, Wu SS (1998) Quantitative assay by flow cytometry of the mitochondrial membrane potential in intact cells. Biochim Biophys Acta 1404: 393-404.

Takahashi T, Broecker WS, Langer S (1985) Redfield ratio based on chemical data from isopycnal surfaces. J Geophys Res 90: 6907-6924.