Vol. 56, No. 7APPLIED AND ENVIRONMENTAL MICROBIOLOGY, JUIY 1990, p. 2234-22420099-2240/90/072234-09$02.00/0Copyright C) 1990, American Society for Microbiology

Rapid Quantification of Planktonic Ciliates: Comparison of ImprovedLive Counting with Other Methods

TELESPHORE SIME-NGANDO,* HANS JULIAN HARTMANN,t AND CLAUDE ALAIN GROLIERE

Laboratoire de Zoologie et Protistologie, Universite Blaise Pascal de Clermont-Ferrand,UA Centre National de la Recherche Scientifique 138, 63177 Aubiere Cedex, France

Received 18 October 1989/Accepted 10 April 1990

The following efficient and quantitatively valid method to filter concentrate and count live planktonic ciliateswas developed and compared with other treatments: unconcentrated (raw) samples and centrifuged sampleswere counted live, and the effects of five different fixatives (HgCl2, Lugol's iodine, formaldehyde, glutaralde-hyde, and Champy-DaFano) on the counts were monitored. Samples originated from a eutrophic mountainlake (Lake Aydat, near Clermont-Ferrand, France). Overall, live filtered counts were similar to counts of rawsamples, but they were significantly higher (2 to 2.3 fold, P < 0.05) by analysis of variance than counts fromcentrifuged samples. Nevertheless, some taxa, i.e., Halteria and Loxodes spp., were sensitive to filtration. Thelive filtered counts were also comparable to counts of raw HgCl2-fixed and settled samples. HgCl2 and Lugolfixation consistently gave the highest total counts, while significantly lower counts were always obtained withChampy-DaFano-fixed samples. Losses due to fixation were insignificant for raw samples but were substantialand statistically significant in concentrated samples (15% after filtration and 71% after centrifugation,compared with counts from the corresponding live samples). Live counting of passively filter-concentratedciliates has many advantages over other methods. It is two to four times quicker and more efficient. Ciliates arerecognized with certainty, more species are identified, and enumeration of dead organisms (e.g., tintinnidloricas) is avoided. It should be recommended as a quantitatively valid alternative to classical methods forassessing planktonic ciliate populations.

Ciliated protozoa are now recognized as important com-ponents of aquatic ecosystems (2, 36, 37, 41). However,knowledge of functional and taxonomic characteristics offield populations frequently is very incomplete (21, 32). Theorganisms are fragile and relatively small, making identifica-tion and counting difficult (14). Quantitative counts areusually made with preserved material under an invertedmicroscope, by using either formaldehyde (27), HgCl2 (34),or Lugol's iodine (23) as a fixative. The preservation invari-ably leads to total or partial destruction of ciliates, limitinginformation needed for identification, especially in sampleswith high detritus content.

Frequently, ciliates are grouped by size (19, 22, 43) todefine their ecological roles. Nevertheless, within a plank-tonic community, ciliate populations may be highly diverse(18, 42), occupying several different trophic compartments(21, 38, 47), regardless of size. Thus, in order to correctlyassign trophic categories (32, 38), routine quantitative ciliatecounts should give well-defined species compositions. Mon-tagnes and Lynn (32) have proposed a routine techniqueimproving the identification of fixed ciliates (quantitativeProtargol stain). However, this technique is elaborate (itrequires 26 steps) and costly and can realistically be usedonly for occasional checks.

Counting live material would be a feasible alternative.Species identification is improved, partially by discriminat-ing between the modes of locomotion of the cells (14). It hasbeen used for counting highly concentrated samples, such asthose found in ponds and in the rumen (15, 18, 20, 24, 39).However, it has rarely been used for counting planktonic

* Corresponding author.t Present address: School of Fisheries, HF 15, Fisheries Teaching

and Research Building, University of Washington, Seattle, WA98195.

ciliates (14, 45, 46). Planktonic samples must usually beconcentrated before statistically valid counts can be ob-tained; this induces stress and a potential loss of cells.We have developed an efficient and quantitatively valid

method to filter concentrate and count live planktonic cili-ates. In this paper, we present tests of its validity byassessing ciliate concentration and species composition fol-lowing several treatments (unconcentrated, filtered, andcentrifuged samples) and comparing live counts with countsof subsamples preserved in five different fixatives.

MATERIALS AND METHODSSite and sampling. Samples were taken with a 5-liter Van

Dorn bottle in Lake Aydat, a small eutrophic lake in theFrench Massif Central (surface area, 0.060 kM2; Zmax,15.5 m) (1; G. Millerioux, Master's thesis, Universitd deClermont-Ferrand II [Universite Blaise Pascal], Aubiere,France, 1976). Samples were taken on three different dates:two in the autumn of 1988 (9 November in the metalimnionat 8 and 10 m and 15 November in the hypolimnion at 13.5 m)and one in the spring of 1989 (5 April in the epilimnion at 4m). These depths corresponded to maximum concentrationsof ciliates in the water column at those dates (Sime-Ngando,unpublished data). The samples were prefiltered through a400-,um-pore-size nylon mesh to eliminate predatory zoop-lankton and stored in dark insulated plastic bottles. In thelaboratory, they were kept in the dark at 4 to 6°C before use.Samples were processed as soon as possible after sampling.Sample processing. The samples collected in autumn were

used only for live counts. The sample collected in spring wasused for both live and preserved counting. Figure 1 summa-rizes the subsampling procedure. The sample was separatedinto three parts: the control subsample (Fig. 1A) (for livecounting of unconcentrated ciliates [Al] and for immediatefixing and later Utermohl counting [A2]) and two treatment

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UNCONCENTRATEDSAMPLES

(CONTROLS)

CONCENTRATEDSAMPLES(TREATMENTS)

Al. UVECOUNTING FILTRATION (1500ml/60ml)(5 or 15 ml)

Bi. Live counting (lml)

1B2. Fixation (1OmI x 5, as A2)

LAKE WATER

SAMPLE

(3600ml)

A2- FIXATION (lOOmI each)- HgCL2 CENTRIFUGATION (1 500ml/60ml)LUGOLS

- Cl. Live counting (imi)

-GLUTARALDEHYDE - C2. Fixation (lOmi x 5, as A2)- CHAMPY - DA FANO

+UTERMOHL COUNTING

FIG. 1. Lake water sample partitioning and treatments of thespring sample.

subsamples concentrated by filtration (Fig. 1B) and centrif-ugation (Fig. 1C) of live material. Following each treatment,samples were either counted live immediately or fixed andcounted later.For filter concentration, six 250-ml subsamples were con-

densed to 10 ml over an 8-jim-pore-size membrane filter(type SC, 45-mm diameter; Millipore Corp.) by using gravityflow or in rare cases a very gentle (<1 kPa) vacuum.Concentration took approximately 20 min. Millipore mem-brane filters have been used with some success to study andcount ciliates in cultures (17), testate amoebae (11, 12, 29),and soil ciliates, i.e., Colpoda aspera (13). Our minimalfiltration vacuum was below values (3 kPa) shown to lead tosignificant losses (15 to 40%) of nanoflagellates (Monas sp.and Bodo sp.) during filtration (8). For centrifugation, livesamples were concentrated by the same factor (25:1) in aseries of 30-ml subsamples with a swinging-bucket refriger-ated rotor (1,200 to 1,300 rpm for 5 min).One portion each of the control and concentrated samples

was retained over ice for immediate live counting. The fiveremaining portions were fixed immediately in one of thefollowing five different preservatives: HgCl2 (2.5% [wt/vol])(34), Lugol's iodine (1% [vol/vol] of the stock solution) (35),formaldehyde (2% [vol/vol] from a 37% commercial solu-tion), glutaraldehyde (0.5% [vol/vol]), and a double fixationof Champy fixative (50% [vol/vol]) followed by centrifuga-tion (two to three times) and conservation in DaFano (9).

Counting. Live counts were done under a stereo micro-scope (magnification, x25 to x50) with a Dolfuss chamber(N. Lair, Ph.D. thesis, Universite de Clermont-Ferrand II

[Universite Blaise Pascal], Aubiere, France, 1975). Thechamber is a shallow rectangular recipient (100 by 200 mm)divided into 200 cubicles (5 by 5 mm) separated by slanted5-mm-deep walls holding about 25 jil of liquid each. Thecubicles are sufficiently small to contain the ciliates forobservation under one or two microscopic fields with rela-tively minor focus adjustments. Temperature fluctuationsare dampened by the heavy-walled glass and by refrigerationprior to use. For counting, 1-ml (concentrated samples) and5- to 15-ml (controls) subsamples were counted by repeat-edly distributing 0.20-ml portions into approximately 10cubicles, to avoid stress due to confinement and exposure tobright light (14). One 1-ml count took 30 to 70 min, and a 5-mlcount took 80 to 120 min.

Fixed samples were counted under an inverted micro-scope by the Utermohl (49) method. For controls, 25-mlportions of the unconcentrated samples were settled for 18 h;for treatments, 1-ml portions of the concentrated sampleswere mixed with 2 ml of filtered lake water (to entirely fill thebase of the 3-ml settling chamber) and settled for 1 h. Ciliateswere counted with a 20x objective, by scanning one half ofthe settling field. One count took at least 90 min. Counting ofpreserved samples was completed within 3 weeks of initialfixation. It was logistically impossible to shorten this time.

All subsample counts (live or preserved) were done intriplicate and occasionally (for live samples collected inautumn) in five replicates. Unknown species and generawere identified by silver staining (48) and/or by the methodsof Kahl (28) and Dragesco and Dragesco-Kerneis (16).

Statistical tests. The counts were tested by analysis ofvariance (ANOVA) for differences between treatments (non-concentrated material versus material with two differentconcentration treatments) and fixatives (live material versusmaterial treated with five different fixatives, and each fixa-tive versus the others). The null hypothesis was as follows:there was no difference between treatments or betweenfixatives or both. Overall significant effects and interactioneffects were tested with two-way ANOVA. Effects withineach treatment category were then further separated byusing one-way ANOVA with a posteriori tests, i.e., theFisher least significant difference test (44) and the Scheffe Ftest (50). Similar tests (ANOVA with a posteriori tests) wereused to assess differences in species composition (number ofspecies found) between treatments and fixatives. All analy-ses were performed with log-transformed data.

RESULTS

Cell counts: global comparisons. A total of 30 taxa werecommonly seen (99%) and used for the count comparison, 26from the autumn samples and 17 from the spring samples.The scuticociliates were as follows (the greatest averagelinear dimensions of 20 to 30 preserved individuals are inparentheses): Cyclidium sp. (15 to 30 ,um; 5 April 1989 and 9November 1988), Dexiotricha plagia (30 to 50 ILm; 5 April1989 and 15 November 1988), Histiobalantium natans (42,um; 9 and 15 November 1988), Pleuronema coronatum (75txm; 9 and 15 November 1988), and Uronema nigricans (40,um; 5 April 1989 and 9 November 1988).The oligotrichs were as follows (the greatest average linear

dimensions of 20 to 30 preserved individuals are in paren-theses): Codonella cratera (64 p.m; 5 April 1989), Halteriasp. (18 ,um; 9 November 1988), Halteria sp. (30 ,um; 9November 1988), Strombidium sp., (28 ,um; 5 April 1989 and9 November 1988), Strombidium viride (45 ,um; 5 April 1989and 9 November 1988), Strombidium sp. (70 ,um; 5 April1989 and 9 November 1988), Strobilidium sp. (21 tLm; 5 April1989 and 9 November 1988), Strobilidium gyrans (45 to 70pum; 5 April 1989 and 9 November 1988), Tintinnidiumfluviatile (90 to 140 ,um; 5 April 1989), and Tintinnopsislacustris (45 to 68 pLm; 5 April 1989).The other taxa were as follows (the greatest average linear

dimensions of 20 to 30 preserved individuals are in paren-theses): Askenasia sp. (30 ,um; 5 April 1989 and 15 Novem-ber 1988), Askenasia volvox (50 ,um; 5 April 1989 and 9November 1988), Coleps hirtus (64 ,um; 9 November 1988),Frontonia sp. (110 ,um; 15 November 1988), Frontonialeucas (230 ,im; 9 and 15 November 1988), Lacrymariapupula (90 ,um; 15 November 1988), Lagynophrya rostrata(65 to 75 ,im; 5 April 1989), Loxodes striatus (124 Jim; 9

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2236 SIME-NGANDO ET AL.

November 1988), Loxodes magnus (200 to 238 ,um; 9 No-vember 1988), Plagiopyla nasuta (80 to 120 ,um; 15 Novem- Eber 1988), Spirostomum teres (109 p,m; 9 November 1988), I +1 +1Spirostomum intermedium (120 to 160 ,um; 9 November r co1988), Urotricha saprophila (30 to 40 ,um; 5 April 1989 and 9November 1988), Vorticella sp. (32 ,um; 5 April 1989 and 15November 1988), and Vorticella sp. (56 ,um; 5 April 1989 and

(0 00 CD9 November 1988). 00 ~ NMeans and standard deviations of total ciliate concentra- +t +1 +1

tions for each treatment-fixative combination are summa- o 0 00

rized in Table 1. For comparisons, we defined the following - -_

variables: T, concentration of the controls (direct counts oflive and preserved samples); E1 and E2, concentration offilter-concentrated and centrifuged samples (live plus pre-served), respectively; L1, L2, etc., index of underestimation E +1+1+or loss rate, i.e., L (percent) = -(100EIT - 100) where E1, 00E2, etc., are to be substituted for E; and R, X, and n, range, e44^nmean, and number of items, respectively.

Variations due to counting among replicates were greaterofor preserved than for live samples. The coefficients of = v CD

variation (CVs) of all the preserved counts (calculated from +I +I +ITable 1) averaged Xcv = 17% (4.6% < R < 30.6%; n = 15), 408while the CVs of live counts averaged onlyXcv = 10%, with _a much narrower range (2.5% < R < 19.4%; n = 11). u

Ciliate concentrations of filtered samples, regardless of oU,

fixation, were relatively close to those of control counts: 0 .. en

XE1/T = 0.87 (0.59 < R < 1.64; n = 9) or a mean loss rate (L1 3.c0 -m= 13%) due to filtration, with individual Els ranging both . +1 +1above and below the control values T (-46% < RL1 < 41%). +1 0 eBy contrast, centrifugation caused a relatively great loss, L2 0._O

. Im

= 54% (16% < RL < 78%; n = 8), and all individual E2swere below T. Champy-DaFano was the most destructivefixative; its counts were lowest in all treatments (Table 1). 4,)oThe differences are confirmed by ANOVA. The two-way X .

ANOVA yielded highly significant effects due to concentra- +1+1+1+tion as well as type of fixative (Table 2), regardless of cell OE E 0

type (i.e., all organisms, scuticociliates, or others dominated ._by oligotrichs). There were also significant interactions, + _especially among the scuticociliates, due to crossover effects 0

eof glutaraldehyde. Q o 0The a posteriori tests further confirmed these observa- o -

tions. They indicated that (i) highly significant differences (P 0 +1 +1 +1< 0.01) were due to centrifugation, especially among scuti- (3 t 0

cociliates (Fig. 2) and (ii) significant to highly significant . . .

differences (P < 0.05 and 0.01) were due to fixation withChampy-DaFano, again particularly among scuticociliates(Fig. 3). 0 CD

Live counts: effects of concentration treatment. We defined eT, as concentration of the live controls (direct counts), E3 u +1 +1 +1and E4 as concentrations of live filter-concentrated and 0Oo _centrifuged samples, respectively, and L3 and L4 as meanloss rates.On the average, the lowest live counts were obtained for

centrifuged samples, with mean losses due to filtration (L3 =15%) and centrifugation (L4 = 28%). However, a detailed N 00look reveals cases in which centrifugation performed at least 2 +1+1+as well as filtration (Fig. 4B and C). r- eo

In the samples collected in the spring, (Fig. 4A), the ioflCflsignificant differences (P < 0.05 and 0.01, respectively) T 0

=

between E4 and E3 as well as T, were largely due to thescuticociliates, while the other groups were less affected by . (e CLcentrifugation. The only differences due to filtration ap- ° o opeared in the samples collected in the fall (Fig. 4B and C). o o EDEvidently, Haltena spp. (L3 = 75%) (Fig. 4B) in the metal- .imnion and large ciliates, primarily Loxodes spp. in the zhypolimnion (L3 = 59%) (Fig. 4C), were sensitive to filtra- m v u

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LIVE COUNTING AND FIXATION OF PLANKTONIC CILIATES 2237

TABLE 2. Two-way ANOVA results for effects of type ofconcentration and fixative on ciliate countsa

ANOVA values for:

All Scutico- OligotrichsMethod organisms ciliates and others

F P F P F P

Concentration (A) 212 0.0001 251 0.0001 40 0.0001Fixation (B) 38 0.0001 49 0.0001 7.3 0.0001Interaction (AB) 5.6 0.0001 7.7 0.0001 2.4 0.03

a Data are from the sample obtained on 5 April 1989. Degrees of freedomare 2, 5, 10, and 36 for A, B, AB, and error, respectively. F, F ratio; P,probability; A, concentration; B, fixative.

tion. When we eliminated these filtration-sensitive taxa fromthe analysis, we obtained, for the metalimnion, L3 = -40%and L4 = 8%, i.e., significantly enhanced counts due tofiltration (Fig. 4B, bars labeled others) and no difference dueto centrifugation. Analogous results were obtained fromsamples from the hypolimnion (Fig. 4C, bars labeled others).

Preserved counts: effects of concentration treatment. Wedefined Tf as the concentration of the preserved controls(direct counts), E5 and E6 as the concentrations of preservedfilter-concentrated and centrifuged samples, respectively,and L5 and L6 as mean loss rates.Loss due to fixation after filtration for all ciliates combined

was relatively small and comparable to that for live counts:Ls = 11%. By contrast, fixation loss after centrifugation wasmuch greater than for live counts, with L6 = 76%, primarilydue to the scuticociliates (Fig. 4A and 5). Similar to livesamples, differences between preserved centrifuged, fil-tered, and direct-counted samples were highly significant.However, in contrast to live samples, preserved oligotrichswere significantly (P < 0.05) reduced in both filter-concen-trated and centrifuged samples, with L5 = 27% (Fig. 5, thirdset of bars).

Effects of type of fixative. Since the HgCI2 counts were onthe average the highest of all fixed counts (Fig. 3), we usedthem as reference counts (H) to compare fixative effects forall treatments combined. By analogy, we defined combined

TOT SCU OL & OTH

ANOVA FP OF FP OF FP SF

16000 DIR vs FOtRS NS NS NO NSDIR vs CEN S . .NS

14000 fitVs CEN NS*O.....,

12000 -

10000 I

a

8000

u6000-

* CONTROL (DIR)

0 FILTRATION (FIL)

El CENTRIFUGATION (CEN)

TOTAL SCUCOMCILIATES OLGOTR"IHS &

FIG. 2. Comparison of effects of concentration treatments on

mean ciliate abundance (cells per liter), live and fixed countstogether. DIR, Direct counts: FIL, counts after filtration; CEN,counts after centrifugation. Data are from the sample obtained on 5April 1989 only. The number of replicates was 3. The inserted tableshows significant differences from one-way ANOVA a posterioricomparisons of treatments (log-transformed data). TOT, Total;SCU, scuticociliates; OLI & OTH, oligotrichs and others; FP,Fisher least significant difference test; SF, Scheffe F test (*, 0.01 <P < 0.05; **, P < 0.01); NS, not significant.

TOT SCU OLI & OTH

ANOVA FP SF FP SF FP SF

LIVE as CHA. * :* * NSHpC2 v CHA. uNNS NOLtWOL Vs CHA. I NSO NS NS NSFOM. vs CPA. * NSO NS NS NOGLUT. vsCHA. * * NSO * NSO * NS

E TOTAL CILIATES

0 SCUTICOCLIATES

*I OUGOTRICHS & OTHERS

LPJE HgCL I . OWTA CHN FA

FIG. 3. Comparison of effects of different fixatives on meanciliate abundance (cells per liter) for all concentration treatments(i.e., control, filtration, and centrifugation) combined. Data are fromthe sample from 5 April 1989 only. Abbreviations and symbols in theinserted table are as defined in the legend to Fig. 2. The number ofreplicates was 3. CHA. and CH Da FA, Champy-DaFano; FORM.,formaldehyde; GLUT and GLUTA, glutaraldehyde.

counts of glutaraldehyde (G), Lugol (1), formaldehyde (F),and Champy-DaFano (C).Champy-DaFano fixation had the greatest loss, compared

with HgCl2 (LC = 61%). It was by far the poorest-qualityfixative. For the remaining fixatives, the overall qualityincreased from formaldehyde to Lugol to glutaraldehyde,i.e., LF = 25%, L, = 17%, and LG = 6%. Differencesbetween Champy-DaFano and Lugol or formaldehyde weremoderately significant (P < 0.05) for scuticociliates, butdifferences between Champy-DaFano and HgCl2 or glutaral-dehyde were always highly significant (P < 0.01) (Fig. 3,inserted table). Thus, glutaraldehyde and HgCl2 are high-quality fixatives, and Lugol and formaldehyde are interme-diate-quality fixatives with inconsistent behavior closer tothat of the poor-quality Champy-DaFano.Combined effect of concentration treatment and fixative.

Significant interactions in two-way ANOVA (Table 2) indi-cated that some fixatives had effects on the concentrationtreatment that were opposite those of others. This appar-ently was due to filter-concentrated scuticociliates fixed inglutaraldehyde (Fig. 6). Filtered and glutaraldehyde-fixedscuticociliates exceeded both HgCl2 and live counts (L5G =-64% and LE5G,E3 = -5%) (cf. Fig. 6). The significantinteractions for total ciliates (F = 4.1, P = 0.02) and forscuticociliates (F = 7.0 and P = 0.0001 for df = 8) disap-peared when glutaraldehyde was excluded from two-wayANOVAs (F = 0.8, P = 0.6 and F = 1.2, P = 0.3,respectively, for df = 6).

Sensitivity of ciliates to fixation following concentration. Wehave already shown that the concentration treatments of livecells caused, on the average, sometimes significant losses ofciliates, i.e., L3 = 15% and L4 = 28% (cf. Fig. 4). We alsofound variable and significant losses of cells due to theaddition of different fixatives, indicating differential sensitiv-ity of live ciliates to fixation (cf. Fig. 3). Thus, regardless ofconcentration treatment, the loss rates in relation to livecounts averaged 7, 15, 23, 31, and 63% for HgCl2, glutaral-dehyde, Lugol, formaldehyde, and Champy-DaFano, re-spectively (grand mean = 28%) (Table 3).When separating treatments, we found that both filter-

concentrated and centrifuged ciliates were much more sen-

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2238 SIME-NGANDO ET AL. APPL. ENVIRON. MICROBIOL.

TOT scU 01 a OTH

ANVA FP SF fP SF FP SFTo 5CU CtlIb OTH

ANOA FP SY fP Sf FP SF

DINoOKNMS MS Mts MS MSl MS20000 *. -M*-*-NS Ms

F. vs0*-CE - " NS18000-

16000

14000-

12000-c

10000'

8000 I

6000

4000-

2000

TOTAL SCJTOLIATOS OUGOTRIKOS&OTHERS

A

ONTROL (DIR)

ILTRATION (FIL)

ENTRFUGATION (CEN)

DURvsFL NS NS WE NSDIRvM CEN

16000 FIL vs CEN

14000

12000

10000C

8000 0 FILTI

6000-M1 CE

4000

2000

TOT MALT TO-MTANOVA M S fP MA FP SF

3600 OMR Fit- "seCEZ *@ ......Vs C N

-fIL,s CEN us

3000

2500

O 2000

1000

0

TOTAL HALTERIA OHllEl

TCT tAR6E OTffRS

*NOVA P sr fP Y FP F

_nDI" fIt 45 NS * * * *3000 -'MOCON * MS

2500

2000c

1500-1

1000

0

TOTAL LAFIG OTHEORS

B

XNTROL (DIR)

:ILTRATION (FIL)

,ENTRFUGATION(CEN)

C

CONTROL (DR)

FILTRATION (FIL)

CENTRIFUGATION (CEN)

TOTAL SCUTrCOCLIATES OUGOTRICHS &OERS

FIG. 5. Comparison of effects of concentration treatments onmean ciliate abundance (cells per liter), fixed samples only. Data arefrom the sample from 5 April 1989 only. Abbreviations and symbolsin the inserted table are as defined in the legend to Fig. 2. Thenumber of replicates was 3.

found per sample is summarized in Table 4. The CVs rangedfrom 0 to 1.8%, indicating little variation among replicates.On the average, more taxa were retained in filter-concen-trated samples (Xfi, = 14.2) than in centrifuged samples(Xcent = 13.1) and controls (Xc.0t = 13.2), essentially be-cause of live counts (Table 4). The live count enhancementdue to filtration was significant in the 8-m sample obtained inthe fall (Xfilt = 20 versus Xcent = 14 and Xc0nt = 14; P < 0.01from one-way ANOVA).By contrast, when comparing fixed samples only, the

number of taxa was equal to that of controls (filtration) orwas reduced (centrifugation). Formaldehyde was destructivein both treatments (P < 0.01 if compared with live samplesand samples treated with HgCl2 and glutaraldehyde), whilethe other fixatives reduced the number of taxa only slightly(no significant differences). In particular, we noted thatAskenasia volvox and Strombidium spp. were sensitive toformaldehyde fixation.

FIG. 4. Comparison of effects of concentration treatments onmean ciliate abundance (cells per liter), live counts only. (A) Datafrom 5 April 1989. (B) Data from November 1988, 8 and 10 m. (C)Data from November 1988, 13.5 m. Large ciliates are primarilyLoxodes spp. Abbreviations and symbols in the inserted table are asdefined in the legend to Fig. 2 (0, no data); number of replicates =3 or 5.

sitive to fixation than the ciliates of controls (Table 3). In thecontrols, there were no losses on the average (XL = -8%),because of gains with HgCl2 (L = -51%) and Lugol (L =-23.3%), and there were substantial losses due to onlyChampy-DaFano (L = 43.5%). Filter-concentrated ciliateswere somewhat sensitive to fixation, with losses averaging15%, similar to live losses. Finally, centrifuged ciliates werehighly sensitive to all fixatives, with an average loss rate of71%, ranging from 60.6% for glutaraldehyde to 85% forChampy-DaFano.

Clearly, the concentration procedures stressed the livingcells and reduced their resistance to most fixatives. Centrif-ugation appeared to be particularly stressful, and there wasno single fixative that performed well under all conditionsand for all types of ciliates.

Effects on species composition. The mean number of taxa

DISCUSSION

The following methods have been proposed for estimatingthe abundance of free-living ciliates: counting preservedsamples in a settling chamber (25), direct counting (3, 13,26), epifluorescence microscopy (40), direct counting of livecells (14), cytobucket quantitative Protargol staining (D. C.

16000'

14000

12000 ,

X 10000'

O 8000'

6000 -

4000'

2000-

* OONL1e] FILTRATION

O3 CENTRFUGATION

LNE HgCL2 LUL FROM GWTA CH Da FA

FIG. 6. Comparison of effects of different concentration treat-ments and fixatives on mean abundance of scuticociliates (cells perliter). Data are from the sample from 5 April 1989; number ofreplicates = 3. Abbreviations are as defined in the legend to Fig. 2.

TROL (DIR)

TRATION (FIL)

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LIVE COUNTING AND FIXATION OF PLANKTONIC CILIATES 2239

TABLE 3. Summary of loss rates obtained from differentconcentration treatments and fixatives compared

with live direct counts

Loss rateTreatment Control Filtra- Centrifu-

(direct counts) tion gation Mean

HgCl2 -51.0 10.4 64.8 7Lugol -23.3 20.9 73.2 23Formaldehyde 6.0 15.6 72.2 31Glutaraldehyde 17.2 -36.1 60.6 15Champy-DaFano 43.5 63.0 85.0 63Mean (all fixatives) -8 15 71 28Live 15 28

Brownlee, Ph.D. thesis, University of Maryland, CollegePark, 1982), and quantitative Protargol staining (32). Directlive counts of unconcentrated samples (3, 14, 45) were thereference standard in most studies comparing quantitativeciliate counting.However, there are a number of critical disadvantages to

direct live counts of unconcentrated samples. In pelagicenvironments, especially in oligotrophic waters, ciliates arerelatively small (10 to 30 p.m) (6, 30) and not very concen-trated (e.g., 1 to 10 cells per ml) (30, 45). Thus, to obtainstatistically valid counts of unconcentrated live samples, agreat volume of water must be scanned (at least 5 ml and upto 100 ml). This is time-consuming (on the average, 4 h forcounting 15 ml of our samples containing 5 to 15 cells per mlwith various-sized ciliates) and therefore cannot be recom-

mended as a routine procedure. Small cells may escapedetection when they are rare or when samples are notimnmediately processed because of a loss of motility. Forexample, we have noted that some of the small oligotrichs,e.g., Strombidium spp. and Strobilidium spp. (10 to 45 p.m inlength), lose motility once exposed for more than 10 minunder microscope light, regardless of temperature.The only alternative to direct live counting is to use

preserved samples or else to concentrate the live samplesbefore counting. Fixation and counting of preserved samplesare currently chosen by most investigators, primarily byusing a settling chamber, and occasionally by epifluores-cence microscopy, cytobucket quantitative Protargol stain-ing, or quantitative Protargol staining. While it does notrequire immediate sample processing, there remain a num-ber of considerable problems with counting fixed samples.Most of our comments relate to the settling chamber.Sample processing times and counting times are relatively

long, even compared with direct live counting; our invertedmicroscope counts took at least 90 min per subsample, andother workers mentioned up to 4 h per sample (F. Rassoul-zadegan, Ph.D. thesis, Universitd de Paris VI [Pierre etMarie Curie], Paris, France, 1982). This disregards prepro-cessing time (10 to 20 min per sample) and limits imposed bythe number of sedimentation chambers available for accom-modating long settling times (e.g., 3 h per cm) (31). Theseconstraints are significant factors when designing a routinecounting program (51). Other techniques (cytobucket quan-titative Protargol staining and quantitative Protargol stain-ing) which allow a more detailed examination of cytologicalstructures and enhance species identification are very elab-orate and costly and in our judgment are not suitable forroutine analyses.

Preserved samples have greater subsampling errors thanlive samples. Our replicate counts of preserved samples hadconsiderably higher CVs (mean CV = 17%) than the livecounts (mean CV = 10%), despite extreme care taken tohomogenize samples before subsampling and to maintainconsistent subsampling procedures for all samples. Thedifference in subsample variability must be due to insuffi-cient homogenization of cells in the preserved samples andnonrandom distribution of cells on the bottoms of the settlingchambers.While the settling method nevertheless may give quanti-

tative information comparable to that of direct live counts,our results clearly demonstrate that its success depends onthe type of fixative used for preservation. Immediate fixationof untreated samples with HgCl2 and Lugol gave comparableor better total numbers than direct live counts. Conversely,Lugol caused relatively great losses in preconcentratedsamples. Glutaraldehyde performed well on the average butnot equally well for all taxonomic groups. Thus, filteredscuticociliates were better preserved with glutaraldehydethan with any other fixative, while untreated or centrifugedones were not (Fig. 6). This could not be a subsamplingerror, because counts of oligotrichs and other cells of any ofthe glutaraldehyde-treated samples (not listed in detail)showed no such inverse treatment-specific differences andthere were no extreme replicate counts (CV of filtered andglutaraldehyde-fixed counts = 12%). Glutaraldehyde clearlywas responsible for the significant interaction between treat-ments and fixatives found by two-way ANOVA (cf. Table 2).Rather, the result illustrates that detrimental fixative effects(i.e., cell disfiguration, rupture, shrinking, etc.) are mini-mized for any given species and fixative at an optimal ratio offixative amount/cell concentration and that an optimal ratio

TABLE 4. Mean number of ciliate taxa identified in the different subsamplesa

Mean no. of ciliate taxa identified in:

Method Spring of 1989 (4 m) Fall of 1988 atb:

Live HgCl2 Lugol Formal- Glutaral- Champy-DaFano 8 m 10 m 13.5 m

Direct counting 14 16 15 15 16 16 14 6 7Filtration 17 16 16 14C 16 16 20d 6e 7Centrifugation 17 16 14 14 15 15 14 6e

a Number of replicates = 3.b All samples were live.c Significant difference (P < 0.01) due to fixations.d Significant difference (P < 0.01) due to treatments.e Number of replicates = 5.s-, No data collected.

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2240 SIME-NGANDO ET AL.

happened to be used for the filtered scuticociliates in ourexperiment.Although total cell counts from two or three of the five

fixatives were roughly comparable to live counts, we notedconsiderable and fixative-specific losses in information forspecies or taxon identification. This has been well docu-mented before (e.g., reference 14). In our samples, 20% ofthe oligotrichs were partially damaged through contact withHgCl2, Lugol, and glutaraldehyde. The damage occurredindependently of filtration or centrifugation. It did not occurin Champy-DaFano-treated samples, where many oligo-trichs may simply have been entirely destroyed upon contactwith the fixatives. We also noted a relatively high CV forAskenasia volvox replicate counts preserved in glutaralde-hyde and formaldehyde, indicating that the cells becomesensitive to subsampling when treated with these fixatives.Since oligotrichs form an important part of planktonic ciliatecommunities (4, 5), fixative-specific misidentification andloss through fixation or subsampling may lead to consider-able underestimation of their abundance and biomass.Other fixative-specific anomalies were noted. Thus, spher-

ical cells, scuticociliates in particular, were easily deformedby Lugol, while the cell contours and the ciliature remainedrecognizable. Cells were less deformed by formaldehyde(10), but cell contours and ciliae became less visible and thecytoplasm clarified except for the vacuoles. In Champy-DaFano, the cytoplasm frequently appeared dark, perhapsbecause of crystallization of one of the components ofChampy; the ciliae disappeared entirely, became bundled(facilitating confusion with detritus particles), or were irreg-ularly distributed around a cell.

Nevertheless, more species-specific information was lostwith formaldehyde than with any other fixative: it was theonly fixative in which significantly fewer taxa were encoun-tered (cf. Table 4). Formaldehyde may be especially damag-ing to a number of structures necessary for identification(e.g., ciliature and cell shape) rather than cause excessivecell ruptures during fixation and handling (34). In addition,since loss rates on total cell counts were relatively great forformaldehyde, we consider it a poor choice for ciliatefixative.Champy-DaFano was by far the worst fixative, for all

types of cells and with all treatments. The loss rates werestaggering and clearly unacceptable for any kind of quanti-tative work. It is possible that the high concentrationsused-50% for Champy and 100% for DaFano, as recom-mended by cytologists (9, 16)-contributed considerably toloss of cells. Losses were enhanced because of the com-pounding effects of two fixatives used in sequence plus theadditional centrifugation required to transfer the Champy-fixed cells to the DaFano solution. This illustrates that anyadditional manipulations of ciliate samples following initialfixation will induce significant cell losses (cf. references 7and 13).

Besides fixative-specific differences, the counting of pre-served samples poses the more general problems of estimat-ing cell numbers and of identification. First, the number oforiginally live tintinnids may be overestimated. In preservedsamples, a tintinnid lorica may be empty (the cell is assumeddead before fixation), filled with a cell and with externallyvisible ciliature (the cell is assumed alive before fixation), orfilled with a retracted cell (ciliae not visible). This last type ofcell may have been dead or alive before fixation. If assumedalive, the number of tintinnids will be overestimated. Con-versely, in live counts, motility helps in determining theviability of tintinnids (14).

Second, optically similar species which may have signifi-cantly different functional adaptations are difficult to distin-guish in preserved samples. Thus, Askenasia volvox is easilyconfounded with Strombidium spp. In general, Strombid-ium, Strobilidium, and Halteria spp. are difficult to separate,especially when their ciliature is hidden by the cell body. Insuch a position they often cannot even be classified withinthe oligotrichs. Sometimes it is difficult to make a distinctionbetween particles and ciliates in general. Obviously, theseproblems are avoided when live material is counted. Suchlimitations usually are not discussed by investigators pub-lishing species lists from studies with preserved material(e.g., references 4 and 33).Our results clearly indicate that concentration by filtration

is preferable to centrifugation. Centrifugation gave signifi-cantly lower counts than controls, both for live samples andfor samples fixed after treatment. Several factors may ex-plain this. The mechanical stress of centrifugation causesrupture of sensitive cells. More importantly, after live cen-trifugation, some ciliates immediately move back into thesupernatant and cannot be recovered through pipetting, evenafter multiple centrifugations. There also appears to be aninherent significant stress associated with centrifugation,more so than with filter concentration. In live counts, theaverage loss rate due to filtration was 15%. In preservedsamples, the average loss rate was also 15% (Table 3) (27.5%when excluding glutaraldehyde, which did not affect scuti-cociliates) (Fig. 6). For centrifuged samples, the two respec-tive rates were higher and the difference between live andfixed losses was great: an increase from 28% (live) to 71%(fixed). The increased loss rate for treated fixed samplescompared with treated live samples (i.e., Table 3, columns 3and 4) is a treatment stress indicator. A corresponding stressindex can be calculated, i.e., D = [1 - (live loss rate)/(fixedloss rate)]. D was lower for filtration (D = 0, or D = 0.45when excluding glutaraldehyde) than for centrifugation (D =0.61), confirming that fixation after filtration was, on aver-age, more benign.We are not aware of any investigators who have centri-

fuged live ciliate samples prior to fixation. Besides savingexpenses for fixatives, there appear to be no inherent advan-tages. Rather, our results show that except for some specialcases (such as cells sensitive to filter contact, in the case ofLoxodes and Halteria spp.), centrifugation of ciliates shouldbe avoided under any circumstances (cf. references 7 and13).A negative effect of filtration on some species has also

been noted by other investigators. Thus, Montagnes andLynn (32) found a filter-sensitive Tetrahymena patula iso-late. Possibly, species react differently to various filtermembranes (e.g., acetate membrane versus polycarbonate),which could be evaluated prior to a routine counting pro-gram. Alternatively, a reverse-pressure filtration techniquecould be tested. Concentration by filtration yielded some-times significantly higher numbers of species than no con-centration or centrifugation (Table 4). This is primarilybecause rare species were not seen or were rarely seen inunconcentrated samples and because live centrifugation mayfavor the loss of particularly motile species (e.g., scuticocil-iates).

In conclusion, we have found that passive filter concen-tration of ciliates together with live counting is statisticallyacceptable for assessing both ciliate abundance and speciescomposition in routine quantitative analyses. Counting andsample processing are relatively rapid (70 to 90 min persample) compared with other methods (up to 4 h per sam-

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LIVE COUNTING AND FIXATION OF PLANKTONIC CILIATES 2241

ple). There are reduced subsampling and counting errors,compared with preserved samples. All the disadvantagesrelated to fixation, with its problems of recognizing anddiscerning many species in a settling chamber, are avoided.Since samples must be processed immediately, there will beno loss due to long-term storage of preserved samples (8;unpublished data). It has been argued that live countingcannot be adapted to shipboard conditions (32). However,the use of a suitable device, such as our Dolfuss chamber,should make counting possible under all extreme oceano-graphic conditions. The method appears well adapted tostudying pelagic ciliates in oligotrophic regions.The two major disadvantages of this method can be

overcome by proper testing and training. The sensitivity ofsome ciliates to the filtration process can be minimized bytesting a variety of filter types and by eliminating filtrationpressure or adapting reverse filtration. Recognition of liveciliate species requires considerable experience; this skillmust be acquired gradually through proper identificationtechniques (16) and with expert advice.

In summary, we found that our method, when carefullycontrolled, is suitable for quantitatively and qualitativelyanalyzing planktonic ciliate populations. It is a statisticallyvalid, practical, rapid, and relatively economical routinemethod. It offers a practical alternative to classical or veryelaborate methods using fixatives.

LITERATURE CITED1. Aleya, L., J. Devaux, H. El Magouri, 0. Marvalin, and C.

Amblard. 1988. Usefulness of simultaneous use of severalmethods for the estimation of phytoplanktonic biomass. Eur. J.Protistol. 23:334-342.

2. Azam, F., T. Fenchel, J. S. Field, L. A. Meyer-Reil, and F.Thingstad. 1983. The ecological role of water-column microbesin the sea. Mar. Ecol. Prog. Ser. 10:257-263.

3. Baldock, B. M. 1986. A method for enumerating protozoa in avariety of freshwater habitats. Microb. Ecol. 12:187-191.

4. Beaver, J. R., and T. L. Crisman. 1982. The trophic response ofciliated protozoans in freshwater lakes. Limnol. Oceanogr.27:246-253.

5. Beaver, J. R., T. L. Crisman, and R. W. Bienert. 1988. Distri-bution of planktonic ciliates in highly coloured subtropicallakes: comparison with clearwater ciliate communities and thecontribution of mixotrophic taxa to total autotrophic biomass.Freshwater Biol. 20:51-60.

6. Beers, J. R., M. H. Reid, and G. L. Steward. 1980. Microplank-ton population structure in southern California nearshore watersin late spring. Mar. Biol. 60:209-226.

7. Berninger, U.-G., B. J. Finlay, and H. M. Canter. 1986. Thespatial distribution and ecology of zoochlorellae-bearing ciliatesin a productive pond. J. Protozool. 33:557-563.

8. Bloem, J., and M.-J. B. Bar Gilissen. 1988. Fixing nanoflagel-lates. Arch. Hydrobiol. Beih. 31:275-280.

9. Chatton, E., and A. Lwoff. 1930. Impregnation, par diffusionargentique, de l'infraciliature des cilies marins et d'eau douce,apres fixation cytoplasmique et sans dessication. C. R. SeancesSoc. Biol. Fil. 104:834-836.

10. Choi, J. W., and D. K. Stoecker. 1989. Effects of fixation on cellvolume of marine planktonic protozoa. Appl. Environ. Micro-biol. 55:1761-1765.

11. Couteaux, M. M. 1967. Une technique d'observation des thec-amoebiens du sol pour l'estimation de leur densite absolue. Rev.Ecol. Biol. Sol 4:593-596.

12. Couteaux, M. M. 1969. Estimation quantitative des thecamoeb-iensedaphiques par rapport a la surface du sol. C. R. Acad. Sci.Ser. III Sci. Vie 281:739-741.

13. Couteaux, M. M., and L. Palka. 1988. A direct counting methodfor soil ciliates. Soil Biol. Biochem. 20:7-10.

14. Dale, T., and P. H. Burkill. 1982. "Live counting"-a quick andsimple technique for enumerating pelagic ciliates. Ann. Inst.

Oceanogr. 58:267-276.15. Darbyshire, J. F. 1972. Nitrogen fixation by Azotobacter chroo-

coccum in the presence of Colpoda steini. I. The influence oftemperature. Soil Biol. Biochem. 4:359-369.

16. Dragesco, J., and A. Dragesco-Kerneis. 1986. Cilids libres del'Afrique intertropicale. Institut Francais de Recherche Scien-tifique pour le Developpement en Cooperation, CollectionFaune Tropicale, 26. Editions de l'ORSTOM, Paris.

17. Erhan, S., E. A. Franko, and R. J. Rutman. 1969. An improvedcounting method for motile protozoan culture. J. Microsc.90:161-164.

18. Fenchel, T. 1969. The ecology of marine microbenthos. IV.Structure and function of the benthic ecosystem, its chemicaland physical factors and the microfauna communities withspecial reference to the ciliated protozoa. Ophelia 6:1-182.

19. Finlay, B. J. 1978. Community production and respiration byciliated protozoa in the benthos of a small eutrophic loch.Freshwater Biol. 8:327-341.

20. Finlay, B. J., P. Bannister, and J. Stewart. 1979. Temporalvariation of benthic ciliates and the application of associationanalysis. Freshwater Biol. 9:45-53.

21. Finlay, B. J., K. J. Clarke, A. J. Cowling, R. M. Hindle, and A.Rogerson. 1988. On the abundance and distribution of protozoaand their food in a productive freshwater pond. Eur. J. Protistol.23:205-217.

22. Gates, M. A. 1984. Quantitative importance of ciliates in theplanktonic biomass of lake ecosystems. Hydrobiologia 108:233-238.

23. Gates, M. A., and U. T. Lewg. 1984. Contribution of ciliatedprotozoa to the planktonic biomass in a series of Ontario lakes:quantitative estimates and dynamical relationships. J. PlanktonRes. 6:443-455.

24. Groliere, C. A., and T. Njine. 1973. Etude comparee de ladynamique des populations de cilies dans differents biotopesd'une mare de foret pendant une annee. Protistologica 9:5-16.

25. Hasle, G. R. 1978. The inverted microscope method, p. 88-96.In A. Sournia (ed.), Phytoplankton manual. United NationsEducational, Scientific & Cultural Organization, Monographs inOceanographic Methodology 6. Page Bros., Ltd., Norwich,United Kingdom.

26. Heal, 0. W. 1964. The use of culture for studying Testacea(Protozoa: Rhizopoda) in soil. Pedobiologia 4:1-17.

27. Hecky, R. E., and H. J. Kling. 1981. The phytoplankton andprotozooplankton of the eutrophic zone of Lake Tanganyika:species composition, biomass, chlorophyll content, and spatio-temporal distribution. Limnol. Oceanogr. 26:548-564.

28. Kahl, A. 1930. Wimpertiere oder Ciliata (Infusoria), p. 1-886. InF. Dahl (ed.), Die Tierwelt Deutschlands, 18 Teil. Urtiere oderProtozoa. Gustav Fischer Verlag, Jena, German DemocraticRepublic.

29. Lousier, J. D., and D. Parkinson. 1981. Evaluation of a mem-brane filter technique to count soil and litter Testacea. Soil Biol.Biochem. 13:209-213.

30. Margalef, R. 1963. R6le des cilids dans le cycle de la viepelagique en Mediterranee. Rapp. Com. Int. Mer Mediterr.17:511-512.

31. Margalef, R. 1969. Counting, p. 7-14. In R. A. Vollenweider(ed.), A manual on methods for measuring primary productionin aquatic environments. F. A. Davis, Philadelphia.

32. Montagnes, D. J. S., and D. H. Lynn. 1987. A quantitativeProtargol stain (QPS) for ciliates: method description and test ofits quantitative nature. Mar. Microb. Food Webs 2:83-93.

33. Pace, M. L. 1982. Planktonic ciliates: their distribution, abun-dance, and relationships to microbial resources in a monomicticlake. Can. J. Fish. Aquat. Sci. 39:1106-1116.

34. Pace, M. L., and J. D. Orcutt. 1981. The relative importance ofprotozoans, rotifers, and crustaceans in a freshwater zooplank-ton community. Limnol. Oceanogr. 26:822-830.

35. Parsons, T., Y. Maita, and C. M. Lalli. 1984. A manual ofchemical and biological methods for seawater analysis. Perga-mon Press, Ltd., Oxford.

36. Pomeroy, L. R., and W. J. Wiebe. 1988. Energetics of microbialfood webs. Hydrobiologia 159:7-18.

VOL. 56, 1990

on May 24, 2018 by guest

http://aem.asm

.org/D

ownloaded from

2242 SIME-NGANDO ET AL.

37. Porter, K. G., M. L. Pace, and J. F. Battey. 1979. Ciliatedprotozoans as links in freshwater planktonic food chains. Na-ture (London) 277:563-565.

38. Pratt, J. R., and J. J. Cairns. 1985. Functional groups in theProtozoa: role in differing ecosystems. J. Protozool. 32:415-423.

39. Senaud, J., C. A. Groliere, B. Zainab, J. Grain, and J. P.Jouany. 1980. Implantation et developpement des populationsde protozoaires cilies (Polyplastron multivesiculatum, Endodi-nium sp., Isotricha prostoma) dans le rumen des moutonsrecevant differents regimes alimentaires. I. Regime Luzerne-Orge (40/50). Protistologica 16:329-337.

40. Sherr, B. F., and E. B. Sherr. 1983. Enumeration of heterotro-phic microprotozoa by epifluorescence microscopy. EstuarineCoastal Shelf Sci. 16:1-7.

41. Sherr, B. F., and E. B. Sherr. 1984. Role of heterotrophicprotozoa in carbon and energy flow in aquatic ecosystems, p.412-423. In M. J. Klug and C. A. Reddy, Current perspectivesin microbial ecology. American Society for Microbiology,Washington, D.C.

42. Small, E. B., and D. H. Lynn. 1985. Phylum Ciliophora, p.393-575. In J. J. Lee, S. H. Hutner, and E. C. Bovee (ed.), Anillustrated guide to the protozoa. Allen Press, Inc., Lawrence,Kans.

43. Smetacek, V. 1981. The annual cycle of protozooplankton in the

APPL. ENVIRON. MICROBIOL.

Kiel Bight. Mar. Biol. 63:1-11.44. Sokal, R. R., and F. J. Rolf. 1981. Biometry. W. H. Freeman

and Co., New York.45. Sorokin, Y. I. 1977. The heterotrophic phase of plankton suc-

cession in the sea of Japan. Mar. Biol. 41:107-117.46. Sorokin, Y. I. 1981. Microheterotrophic organisms in marine

ecosystems, p. 293-342. In A. R. Longhurst (ed.), Analysis ofmarine ecosystems. Academic Press, Inc. (London), Ltd., Lon-don.

47. Stoecker, D. K., and G. T. Evans. 1985. Effects of protozoanherbivory and carnivory in a microplankton food web. Mar.Ecol. Prog. Ser. 25:159-167.

48. Tuffrau, M. 1967. Perfectionnements et pratique de la techniqued'impregnation au protargol des infusoires cilies. Protistologica3:91-98.

49. Utermohl, H. 1958. Zur Vervollkommnung der quantitativenPhytoplankton-Methodik. Mitt. Int. Ver. Theor. Angew. Lim-nol. 9:1-38.

50. Winer, B. J. 1971. Statistical principles in experimental design.McGraw-Hill Book Co., New York.

51. Woelkerling, W. J., R. R. Kowal, and S. B. Gough. 1976.Sedgwick-Rafter cell counts: a procedural analysis. Hydrobio-logia 48:95-107.

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.org/D

ownloaded from