APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Oct. 2001, p. 4896–4900 0099-2240/01/$04.00⫹0 DOI: 10.1128/AEM.67.10.4896–4900.2001 Copyright © 2001, American Society for Microbiology. All Rights Reserved.
Vol. 67, No. 10
Assessing the Diversity of Marine Bacterial -Glucosidases by Capillary Electrophoresis Zymography† ´ S M. ARRIETA JESU
AND
GERHARD J. HERNDL*
Department of Biological Oceanography, Netherlands Institute for Sea Research, 1790 AB Den Burg, Texel, The Netherlands Received 7 May 2001/Accepted 31 July 2001
We propose a new method for the fast separation and detection of -glucosidases in environmental samples. With this approach, -glucosidases extracted from bacteria are evidenced by substrate-incorporated capillary electrophoresis (CE zymography) and their kinetic parameters can be determined by repeated injections using different substrate concentrations. Preliminary results obtained with natural bacterial communities from the coastal North Sea suggest that the diversity of -glucosidases in the marine environment might be much higher than previously observed. not only for ectoenzymatic activity but also for uptake of organic substrates (4, 21, 23). The obvious question which arises from these studies is as follows: why should natural bacteria exhibit Km values in the millimolar range if the substrate is present only in the nanomolar to micromolar range in the environment? Another question is whether a single bacterial species or even an individual cell can exhibit biphasic uptake and ectoenzymatic activity. To the best of our knowledge, there are no reports of bi- or multiphasic hydrolytic ectoenzyme kinetics available for single bacterial strains. The potential ecological implications of these observations have been discussed previously (3, 23). Without going into detail here, the problem of the apparently existing ectoenzymatic diversity in the natural environment has not been addressed yet. There is only one study which determined the diversity of a bacterioplankton ectoenzyme, -glucosidase, in the natural environment (19). These authors found only two different -glucosidases in the pycnocline layer of the Adriatic Sea, indicating a rather limited diversity of -glucosidase, especially if the large number of potential substrates is considered (19). With the present methods available for assessing ectoenzymatic activity under near-natural conditions, the required resolution to determine ectoenzyme diversity in natural bacterioplankton communities cannot be achieved. We therefore modified a capillary electrophoresis (CE)-based method, originally described by Xue and Yeung (26) for detecting lactate dehydrogenase in mammalian cells, to separate and detect the different bacterial -glucosidases present in seawater. This modified method not only allows the determination of the ectoenzyme diversity present in a given sample but also allows the simultaneous determination of the kinetics of all the different ectoenzymes present in the sample. In this paper we present the outline of the method with the example of -glucosidase. (This work was performed in partial fulfillment of the requirements for a Ph.D. from the University of Groningen, Groningen, The Netherlands, by J.M.A.) Bacterial ectoenzyme extraction. For developing this method, four -glucosidase-producing bacterial strains (two gram-positive strains, Planococcus citreus and Salinicoccus roseus, and two gram-negative strains, Vibrio sp. and Paracoccus alkenifer, belonging to the ␥- and ␣-proteobacteria, respec-
Bacterioplankton are the principal consumers of the dissolved organic carbon (DOC) pool in the sea, which besides soil humus represents the largest reactive organic carbon reservoir on earth (24). The chemically characterizable macromolecular compounds of the DOC pool are carbohydrates, proteins, and lipids, comprising all together about 20 to 40% of the total DOC, commonly with higher percentages of macromolecular compounds present in the euphotic layers of the ocean than in the deep waters (5). The turnover of this DOC pool is highly variable, ranging between 40 to 100 days in the euphotic layers and averaging 6,000 years in the deep sea (14, 25). As shown by ultrafiltration techniques, only about 20 to 30% of the DOC present in the ocean is of ⬎1,000 Da (2). Yet this high-molecular-size fraction, which is contemporarily produced (20), is turned over more rapidly than the ⬍1,000-Da DOC fraction (1, 2). Gram-negative bacterioplankton, which usually constitute more than 90% of the total bacterioplankton in seawater (9), can only transport molecules of ⬍600 Da through their cell walls (17). This hydrolysis of the large bioavailable molecules is mediated by surface-bound ectoenzymes located at the cell wall or in the periplasmic space (8, 18). Usually bacterial surface-associated ectoenzymatic activity dominates over freely dissolved extracellular enzymes (sensu Chro ´st [8]) in the marine environment (6). Over the past two decades, a number of studies have focused on the determination of bacterial ectoenzymatic activity using substrate analogs linked to fluorophores which fluoresce upon cleavage from the substrate by the activity of the appropriate ectoenzymes (12, 13). The high fluorescence yield upon cleavage allows determination of the ectoenzymatic activity of bacterioplankton using incubation periods of only minutes to hours and in situ temperature conditions. This approach has been widely applied to determine the ectoenzymatic activity of bacterioplankton. Biphasic or even multiphasic kinetics have been obtained
* Corresponding author. Mailing address: Department of Biological Oceanography, Netherlands Institute for Sea Research (NIOZ), P.O. Box 59, 1790 AB Den Burg, Texel, The Netherlands. Phone: 31-222369-507. Fax: 31-222-319-674. E-mail:
[email protected]. † This is NIOZ contribution number 3631. 4896
VOL. 67, 2001
ECTOENZYME SEPARATION BY CE
4897
TABLE 1. Mean extraction efficiency and range of -glucosidase activity measured in triplicate for the single bacterial strains and in five different bacterial communities from the coastal North Sea Strain or community
Extraction efficiency ⫾ SD (%)
Vibrio sp. P. alkenifer S. roseus P. citreus Natural community
92.8 ⫾ 5.06 53.5 ⫾ 2.74 23.5 ⫾ 1.87 0.3 ⫾ 0.02 73.4 ⫾ 9.7
Range
67.2–83.6
tively) isolated from the northern Adriatic Sea (11) were used. The strains were cultured in ZoBell 2216 broth (5 g of peptone, 1 g of yeast extract, 1 liter of 0.2-m-pore-size-filtered seawater). One-milliliter aliquots of exponentially growing cultures were harvested by centrifugation (3,200 ⫻ g, 4°C, 15 min) and washed three times with 0.2-m-pore-size-filtered artificial seawater (16) before resuspension of the pellet in the buffer solution for ectoenzyme extraction (described below). Natural bacterioplankton communities were collected from the coastal North Sea using acid-rinsed carboys. Fifty liters of seawater was filtered through 0.8-m-pore-size polycarbonate filters (142-mm diameter; Millipore, Bedford, Mass.) in order to exclude most eukaryotic organisms. To minimize the loss of bacterial biomass due to clogging, the filter was replaced every 10 liters. Bacteria in the filtrate were concentrated to a final volume of 0.5 liters using a Pellicon (Millipore) tangential-flow filtration system equipped with a 0.1-m-pore-size filtration cartridge (hydrophilic polyvinylidene difluoride membrane [Durapore]; Millipore). The bacteria in the retentate were further concentrated by centrifugation (20,000 ⫻ g, 4°C, 30 min) and the resulting pellet was washed three times with artificial seawater. About 108 to 109 bacterial cells obtained from bacterial cultures or from natural bacterial communities were resuspended in 1 ml of extraction buffer consisting of 40% glycerol (Sigma, St. Louis, Mo.), 100 mM taurine (Fluka Chemie AG, Buchs, Switzerland), 20 mM cholic acid (Fluka), and 1 mM MgSO4 (pH 7.50), sonicated on ice at 50 W for 30 s, and centrifuged again for 60 min (20,000 ⫻ g, 4°C). Thereafter, the supernatant containing the enzyme extract was carefully siphoned off and stored at 4°C for subsequent analysis. The bulk of the intracellular components such as DNA and RNA remained in the pellet, as verified by phenol-chloroform nucleic acid extraction and agarose gel electrophoresis (data not shown). Therefore, we assume that the enzymes in the supernatant are ectoenzymes. The -glucosidase activity of both the strains and the natural bacterial communities was measured in triplicate subsamples by means of the fluorogenic substrate analog methylumbelliferyl--D-glucoside (13) in the cells resuspended in artificial seawater before extraction, in the enzyme extract, and in the remaining pellet in order to determine the extraction efficiency for -glucosidase. The fluorescence was measured at an excitation wavelength of 360 nm and an emission wavelength of 445 nm on a Hitachi F-2000 spectrofluorometer (Hitachi, Tokyo, Japan). The extraction efficiencies ranged from 0.3% for the gram-positive P. citreus to 92.8% for the Vibrio sp. (Table 1). For the freshly collected natural bacterial communities the
FIG. 1. Schematic outline of the method used to separate and detect different types of -glucosidases. Different symbols represent different enzymes in the sample (see text for a more detailed explanation of the four steps).
average extraction efficiency was 73.4%. Extracting the -glucosidases in the presence of Triton X-100 as previously described (19) did not improve the extraction efficiency. Moreover, the presence of Triton X-100 in the sample interfered with the separation (data not shown). No effort was made to inactivate proteases in the sample since the electrophoretic patterns remained unchanged in shape and relative intensity during storage in the dark at 4°C for 4 days. CE separation and online detection of -glucosidase activity. Analyses were performed using a Biofocus 3000 CE system equipped with a Biofocus LIF2 laser-induced fluorescence detector (Bio-Rad, Hercules, Calif.). The enzyme separation and detection were performed in 50-m-inside-diameter fused silica capillaries. To avoid electro-osmotic flow and to minimize protein interaction with the wall of the capillary, the inner surface of the capillary was coated with polyacrylamide by means of a siloxane bond (15). The total length of the capillary was 75 cm and the distance from the inlet to the detection window was 70.4 cm. Detection of -glucosidase activity in the capillary was based on the hydrolysis of the fluorogenic substrate analog resorufin--D-glucopyranoside (Rglu) (Sigma). The fluorophore released upon hydrolysis is resorufin and was measured using the 594-nm line of the helium-neon laser as the excitation source, with the emission measured at 630 nm in the LIF2 system (600DRLP02 beam splitter, 630DF30 discrimination filter; Bio-Rad). The electrophoresis buffer contained 100 mM taurine, 20 mM cholic acid, and 1 mM MgSO4 dissolved in distilled water, and the pH was adjusted with NaOH to 7.50. Different concentrations of the substrate (Rglu) were added to the electrophoresis buffer, ranging from 0.1 to 350 M final concentrations; however, at concentrations higher than 200 M, a small amount of precipitate was observed after a few hours of storage at 4°C, and those results were excluded from the calculations of enzyme kinetics. The samples containing the solubilized ectoenzymes and buffer solutions were transferred to the refrigerated carousel of the Biofocus system and kept at 4°C. The capillary temperature was set at 15°C to protect the enzymes from Joule heating during the electrophoretic separation using FC-77 (3M, St. Paul, Minn.) as a cooling fluid. The activity of the different -glucosidases was measured using a four-step procedure (Fig. 1) similar to that described by
4898
ARRIETA AND HERNDL
Xue and Yeung (26) to measure lactate dehydrogenase activities in erythrocytes. The solubilized enzymes were hydrodynamically injected into the capillary at a constant pressure ⫻ time value of 15 lb/in2 ⫻ s to provide a fixed injection volume of 147 nl as calculated by repeated injection and pressure mobilization of a resorufin standard. Thereafter, 20 kV was applied for 20 min to separate the different -glucosidases based on their different electrophoretic mobility and to allow them to migrate into the region containing the substrate (Fig. 1). Due to the presence of glycerol in the sample buffer, the ionic strength in the sample zone is lower than in the electrophoresis buffer, producing a transient decrease in the present level during the first 3 min of the separation as observed for all the runs. This phenomenon is known as field amplification and might be responsible for an additional sample concentration and narrowing of the observed peaks (7). The voltage was then turned off to allow the enzymes to hydrolyze the fluorogenic substrate added to the electrophoresis buffer. Hydrolysis of this fluorogenic substrate leads to a local accumulation of the fluorescent hydrolysis product (resorufin) in those regions of the capillary where -glucosidases are present. After allowing the different -glucosidases to react with the Rglu substrate for 10 to 20 min (see below), 20 kV was applied again to elute the fluorescent products passing the detection window where quantification takes place. Separation of different -glucosidases. The separation conditions were optimized for -glucosidase suspensions extracted from two different strains of gram-negative bacteria (Vibrio sp. and P. alkenifer). Single peaks of -glucosidase activity were obtained when the extracts from the two bacterial strains were injected separately (Fig. 2A). The fluorescent signal increased linearly with increasing incubation times of 5, 10, 20, 30, and 40 min, without significant peak distortion due to diffusion of the fluorescent product. However, a tail of elevated fluorescence was found preceding the peaks (Fig. 2A) in all electropherograms. The intensity of this feature was proportional to the amount of sample injected but did not increase with increasing incubation times and did not appear in heat-denatured samples. Therefore, we believe that it is produced by the enzyme(s) during its migration in the capillary. A similar behavior was also reported for lactate dehydrogenase (27). This fluorescent tail, however, biases the determination of an accurate baseline, especially if dealing with more complex samples such as natural enzyme extracts. To accurately determine hydrolysis rates and evidence the enzyme peaks, repeated injections with increasing incubation periods are required in order to discriminate the fluorescence produced during the incubation period from the signal produced elsewhere. Therefore, incubation times of 10 and 20 min were chosen for the experiment with the natural community and the ectoenzymatic activity was calculated by integrating the peak area. Different pH values ranging between 7.50 and 8.50 were tested. The peak resolution decreased at pHs of ⱖ8.00. Therefore, pH 7.50 was chosen for an optimal separation of the -glucosidases. To determine the potential of the CE to separate -glucosidases, equal amounts of the extracts from Vibrio sp. and P. alkenifer were mixed. As shown in Fig. 2B, the mobility of an individual -glucosidase is independent of the presence of other -glucosidases in the same sample and the amount of
APPL. ENVIRON. MICROBIOL.
FIG. 2. Electropherograms obtained after injecting enzyme extracts from two different strains (Vibrio sp. and P. alkenifer) separately (A) and mixed together (B); note that the peak height is reduced by 50% in the mixture (B) as it contains only 50% of each of the extracts. RFU, relative fluorescence units.
fluorescent product released is proportional to the amount of enzyme injected. Separation of -glucosidases from natural bacterial communities. Once the method was optimized for the separation of -glucosidases from strains, the ectoenzymes from a whole bacterial community were extracted as described above and run in the CE under the same conditions described for the strains. Peaks were considered to represent -glucosidases if the integrated area of fluorescence increased systematically in replicate incubations of 10 and 20 min, as explained above. In contrast to previous observations on the variability (diversity) of -glucosidases in marine systems (19), the electrophoretic pattern we obtained (Fig. 3) revealed seven distinct peaks of -glucosidase activity, while Rath and Herndl (19) found only one to two different types of -glucosidase using chromatographic methods. The differences between the former study and the CE approach may be partly due to the superior resolution power of CE in combination with the sensitivity of laserinduced fluorescence detection. The differences in electrophoretic mobility of native proteins are caused by differences in the amino acid sequence and/or by the secondary structure of the protein (10). Therefore, these different forms of -glucosidase may have different functional properties such as reaction kinetics. Differences in the kinetic properties of ectoenzymes often reflect differences in the substrate supply and utilization among different members of the bacterioplankton community (22, 23).
VOL. 67, 2001
ECTOENZYME SEPARATION BY CE
4899
TABLE 2. Kinetic parameters of the different -glucosidases detected in a natural bacterioplankton sample Peak no.a
Vmax (fmol of resorufin min⫺1)
Km (mol liter⫺1)
1 2 3 4 5 6 7
1,687.79 416.84 203.57 328.94 309.01 111.29 288.47
70.09 36.92 79.87 136.78 26.02 37.69 34.58
a
FIG. 3. Electrophoretic pattern obtained from enzymes extracted from a natural bacterial community. The different lines show the results obtained with different concentrations of Rglu. RFU, relative fluorescence units.
Determination of -glucosidase kinetics using CE. Standard solutions of resorufin ranging from 8 nM to 400 M were injected in triplicate to establish the calibration curves necessary for converting the relative fluorescence units into resorufin concentrations. Therefore, the absolute amount of resorufin injected ranged from 0.90 to 58,800 fmol (1 fmol ⫽ 10⫺15 mol). Despite the fact that the 594-nm wavelength provided by our laser is not optimal for the excitation of resorufin (excitation maximum, 571 nm), we successfully detected as few as 3.45 ⫻ 1010 molecules (57.33 fmol) of resorufin in the electrophoresis buffer. Below this threshold it was not possible to clearly distinguish the signal from background noise. The increase in fluorescence intensity of resorufin was linear (r2 ⫽ 0.998) over the whole range from 57.33 up to 58,800 fmol of resorufin, making this method suitable for quantification of enzyme activity over a large range of enzyme concentrations and specific activities. At the lowest amounts of resorufin (⬍1 pM), the measured peak area was slightly lower than expected from the calculated regression line obtained over the whole concentration range. However, this had no effect on our measurements due to the high fluorescent readings obtained for both the strains and the natural samples. The kinetic properties of the detected -glucosidases were determined with triplicate injections of the enzyme solutions in the electrophoresis buffer containing concentrations of the fluorogenic substrate Rglu ranging from 0.05 to 150 M. An example is given in Fig. 3 showing the pattern obtained from repeated injections of the -glucosidases extracted from a natural sample at different substrate concentrations. The hydrolysis rates corresponding to each of the detected peaks were first calculated for each concentration and then used to calculate the specific Vmax and Km values of the different -glucosidases by nonlinear regression (8). The estimated Km values of -glucosidases obtained from coastal North Sea bacterioplankton ranged from 26.02 to 136.78 mol liter⫺1 (Table 2). Characterizing the diversity of -glucosidases. For the characterization of the -glucosidase diversity it is necessary to measure both the number of different -glucosidases present in the environmental sample and the relative amount of each
Peak numbers correspond to the peaks shown in Fig. 3.
ectoenzyme (Vmax is proportional to the amount of enzyme present). The relative amount of ectoenzyme is certainly biased due to the different extraction efficiencies observed. However, if we assume that the extraction efficiency is constant for a specific ectoenzyme produced by a bacterial species, it is possible to compare the relative abundance of a specific -glucosidase between related samples, i.e., by studying the succession of ectoenzyme activities in bacterioplankton communities. Different ectoenzymes might have the same electrophoretic mobility, thus potentially leading to an underestimation of the number of enzymes present. Therefore, the determination of the -glucosidase richness as obtained by CE has to be considered a conservative estimate. Nevertheless, we detected a higher number of -glucosidases in natural bacterioplankton with the CE approach than that reported hitherto. It is now possible to estimate phylogenetic microbial diversity in natural samples using molecular techniques (9). However, the link between the phylogenetic and functional diversity of microbial communities is still largely missing. Despite the above-mentioned limitations of the CE approach, we believe that it is a new and promising tool for determining functional bacterial diversity. Although it is not possible to directly link the different types of -glucosidase to the bacterial species producing them, it is possible to relate the changes in the phylogenetic composition observed in bacterioplankton communities to the shifts in the enzyme composition of the community. The CE approach is not limited to the determination of -glucosidase activity, but by using other substrates it should also be possible to detect the variability of other enzymes occurring in marine environments. Finally, with some modifications in the extraction protocols and buffer system used, the CE approach should be suitable for studying ectoenzyme diversity in other natural environments as well, such as freshwaters and soils. Financial support was provided by the Dutch Earth and Life Sciences Research Council (ALW) and the NIOZ. J.M.A. was supported by a predoctoral grant from the Basque Government. REFERENCES 1. Amon, R. M. W., and R. Benner. 1996. Bacterial utilization of different size classes of dissolved organic matter. Limnol. Oceanogr. 41:41–51. 2. Amon, R. M. W., and R. Benner. 1994. Rapid cycling of high-molecularweight dissolved organic matter in the ocean. Nature 369:549–552. 3. Azam, F., and B. C. Cho. 1987. Bacterial utilization of organic matter in the sea, p. 261–281. In M. Fletcher (ed.), Ecology of microbial communities. Cambridge University Press, Cambridge, United Kingdom. 4. Azam, F., and R. E. Hodson. 1981. Multiphasic kinetics for D-glucose uptake by assemblages of natural marine bacteria. Mar. Ecol. Prog. Ser. 6:213–222. 5. Benner, R., B. Biddanda, B. Black, and M. McCarthy. 1997. Abundance, size distribution, and stable carbon and nitrogen isotopic compositions of marine
4900
ARRIETA AND HERNDL
organic matter isolated by tangential-flow filtration. Mar. Chem. 57:243–263. 6. Bochdansky, A. B., S. Puskaric, and G. J. Herndl. 1995. Influence of zooplankton grazing on free dissolved enzymes in the sea. Mar. Ecol. Prog. Ser. 121:53–63. 7. Burgi, D. S., and R. L. Chien. 1996. Application and limits of sample stacking in capillary electrophoresis, p. 211–226. In K. D. Altria (ed.), Capillary electrophoresis guidebook: principles, operation and applications, vol. 52. Humana Press Inc., Totowa, N.J. 8. Chro ´st, R. J. 1991. Environmental control of the synthesis and activity of aquatic microbial ectoenzymes, p. 29–59. In R. J. Chro ´st (ed.), Microbial enzymes in aquatic environments. Springer-Verlag, New York, N.Y. 9. Giovannoni, S., and M. Rappe´. 2000. Evolution, diversity, and molecular ecology of marine procaryotes, p. 47–84. In D. L. Kirchman (ed.), Microbial ecology of the oceans. John Wiley & Sons, New York, N.Y. 10. Hedrick, J. L., and A. J. Smith. 1968. Size and charge isomer separation and estimation of molecular weights of proteins by disc gel electrophoresis. Arch. Biochem. Biophys. 126:155–164. 11. Herndl, G. J., J. M. Arrieta, and K. Stoderegger. 1999. Interaction between specific hydrological and microbial activity leading to extensive mucilage formation in the northern Adriatic Sea. Ann. Ist. Super. Sanita 35:405–409. 12. Hoppe, H.-G. 1991. Microbial extracellular enzyme activity: a new key parameter in aquatic ecology, p. 60–83. In R. J. Chro ´st (ed.), Microbial enzymes in aquatic environments. Springer Verlag, New York, N.Y. 13. Hoppe, H.-G. 1983. Significance of exoenzymatic activities in the ecology of brackish water: measurements by means of methylumbelliferyl-substrates. Mar. Ecol. Prog. Ser. 11:299–308. 14. Kirchman, D. L., Y. Suzuki, C. Garside, and H. W. Ducklow. 1991. High turnover rates of dissolved organic carbon during a spring phytoplankton bloom. Nature 352:612–614. 15. Li, S. F. Y. 1996. Capillary electrophoresis: principles, practices and applications, vol. 52, p. 155–200. Elsevier, Amsterdam, The Netherlands. 16. Parsons, T., Y. Maita, and C. Lalli. 1984. A manual of chemical and biological methods for seawater analysis. Pergamon Press, Oxford, United Kingdom.
APPL. ENVIRON. MICROBIOL. 17. Payne, J. W. 1980. Transport and utilization of peptides by bacteria, p. 212–256. In J. W. Payne (ed.), Microorganisms and nitrogen sources. John Wiley and Sons, Chichester, United Kingdom. 18. Priest, F. G. 1984. Extracellular enzymes, vol. 9. Van Nostrand Reinhold, Wokingham, United Kingdom. 19. Rath, J., and G. J. Herndl. 1994. Characteristics and diversity of -D-glucosidase (EC 3.2.1.31) activity in marine snow. Appl. Environ. Microbiol. 60:807–813. 20. Santschi, P. H., L. Guo, M. Baskaran, S. Trumbore, J. Southon, T. S. Bianchi, B. Honeyman, and L. Cifuentes. 1995. Isotopic evidence for the contemporary origin of high-molecular-weight organic matter in oceanic environments. Geochim. Cosmochim. Acta 59:625–631. 21. Talbot, V., and M. Bianchi. 1997. Bacterial proteolytic activity in sediments of the subantarctic Indian Ocean sector. Deep-Sea Res. Part II 44:1069– 1084. 22. Tholosan, O., F. Lamy, J. Garcin, T. Polychronaki, and A. Bianchi. 1999. Biphasic extracellular proteolytic enzyme activity in benthic water and sediment in the northwestern Mediterranean Sea. Appl. Environ. Microbiol. 65:1619–1626. 23. Unanue, M., B. Ayo, M. Agis, D. Slezak, G. J. Herndl, and J. Iriberri. 1999. Ectoenzymatic activity and uptake of monomers in marine bacterioplankton described by a biphasic kinetic model. Microb. Ecol. 37:36–48. 24. Williams, P. J. L. 2000. Heterotrophic bacteria and the dynamics of dissolved organic material, p. 153–200. In D. L. Kirchman (ed.), Microbial ecology of the oceans. Wiley-Liss, New York, N.Y. 25. Williams, P. M., and E. R. M. Druffel. 1987. Radiocarbon in dissolved organic matter in the North Pacific Ocean. Nature 330:246–248. 26. Xue, Q., and E. S. Yeung. 1994. Variability of intracellular lactate dehydrogenase isoenzymes in single human erythrocytes. Anal. Chem. 66:1175–1178. 27. Xue, Q. F., and E. S. Yeung. 1996. Determination of lactate dehydrogenase isoenzymes in single lymphocytes from normal and leukemia cell lines. J. Chromatogr. B 677:233–240.
