catenella species complex: Pyridine linked dehydrogenases

BiochemicalSystematicsand Ecology,Vol. 14, No. 3, pp. 311-323, 1986. Printed in GreatBritain.

0305-1978/86$3.00+0.00 PergamonJournals Ltd.

Electrophoretic Variability within the

Protogonyaulax tamarensis/catenella Species Complex:

Pyridine Linked Dehydrogenases

ALLAN D. CEMBELLA and F. J. R. TAYLOR University of British Columbia, Departments of Botany and Oceanography, 6270 University Boulevard, Vancouver, Canada V6T lW5

Key Word Index--Protogonyaulax species; Gonyaulacales; dinoflagellates; isozymes, electrophoresis; dehydrogenases; linkage dendrogram; taxonomy. Abstract--Extracts from 20 isolates of the Protogonyaulax (= Gonyaulax) tamarensis/catenella species complex from diverse geographical locations, including ten contemporaneous isolates from the same geographical population, were subjected to enzyme electrophoresis. Analysis of isozyme banding patterns produced by eight pyridine-linked dehydrogenases revealed a high degree of genetic polymorphism within and between morphotypes and geographical populations. Since there was little apparent correlation between electromorphs and variants assigned provisionally to the catenelloid or tamarensoid morphotype, and because morphological intermediates exist, the presently used morphological characters used to discriminate unequivocally between R catenella and R tamarensis appear to be inadequate. Nevertheless, within a given morphotype, isolates from the same location were more similar than to those from elsewhere. Isozyme electrophoresis offers a means of biochemically discriminating between isolates of this species complex, for which the plate patterns are substantially the same.

Introduction Members of the Protogonyaulax (=Gonyaulax) tamarensis/catenella species complex are toxigenic thecate dinoflagellates, which are the primary cause of paralytic shellfish poisoning (PSP) in temperate coastal waters throughout the world. The taxonomic status of this group has recently undergone several revisions, with the aim of correcting inadvertent errors made in previous treatments and to identify biologically meaningful affinities within the group. Several recent reviewers [1-7] have attempted to resolve these nomenclatural difficulties, basing arguments largely upon morphological features, including general cell form, thecal plate tabulation and shape, the structure of the apical pore complex (APC), the occurrence and orientation of the attachment pore on the posterior sulcal plate, the presence or absence of a ventral notch (pore) on the 1' apical plate, cyst type, and chain length of colonial forms. At this point, a generally accepted consensus has not yet emerged regarding the generic assignation of this species complex, although it is clear that the removal of this group from the genus, Gonyaulax, is war(Revised received 15 May 1985)

311

ranted [2, 5-7]. Within this species complex, we have encountered difficulties in distinguishing between P. tamarensis and P. catenella, which may co-occur or occur adjacently in the North East Pacific coast region. Frequently, morphological intermediates are noted in boundary zones, areas which are geographically intermediate. Braarud [8] noted that clonal isolates referable to P. (= Gonyaulax tamarensis on the basis of identical thecal plate tabulation, which displayed considerable overlap in other morphological features, nevertheless showed identifiable differences in physiological responses. This suggested that genotypic variants may not be morphologically distinguishable and underscored the requirement for the examination of large numbers of cells through repetitive isolation and subculturing to determine intraspecific affinities. The degree of morphological plasticity exhibited by members of this species complex when they are brought into culture necessitates the application of biochemical techniques to clarify taxonomic relationships. Biochemical approaches which have been previously applied to investigate taxonomic affinities within the group have involved comparative studies of

312

toxicity [9-12] and bioluminescence [3, 11]. Gel electrophoresis of proteins is a useful technique which has been frequently used to discriminate between and within assemblages of a wide variety of lower eukaryotic organisms, including protozoa [13-16], macroalgae [17-19], and microalgae [20], particularly diatoms [21-24]. Electrophoretic studies on dinoflagellates are not numerous in the literature: Schoenberg and Trench [25] examined electrophoretic variability and host specificity relationships for the symbiotic dinoflagellate, Symbiodinium (= Gymnodinium) microadriaticum, based upon four enzymes and total soluble protein; Watson and Loeblich [26] used electrophoretic isozyme analysis to infer evolutionary relationships between potential sibling species within the marine genus, Heterocapsa, while Beam et al. [27] applied this technique similarly to the study of Crypthecodinium cohnii, a marine heterotroph. However, with the exception of work by Hayhome and Pfiester [28] on 11 clonal isolates of freshwater Peridinium spp., separation of dinoflagellate isozymes has been performed on starch gels, and has typically involved only a few enzyme systems. In an effort to achieve superior isozyme band resolution and reproducibility, we have used polyacrylamide gels (PAG), rather than starch gels. Staining systems were perfected from standard methods for eight pyridine-linked dehydrogenases, enzymes for which the specific in vivo substrates are known, and which are considered to be relatively genotypically conservative [29, 30]. Soluble enzyme extracts from 20 isolates of the Protogonyaulax tamarensis/catenella complex, brought into culture from diverse geographical regions were subjected to PAG electrophoresis. Correspondence between isozyme banding profiles indicated that contemporaneous isolates from the same location were more closely related to each other, than to isolates obtained from outside the region. Significantly, the isozyme data did not support the establishment of a clear line of demarcation between isolates conforming to the tamarensoid versus the catenelloid morphotype. Results The isozyme banding patterns for the 20 Protogonyaulax isolates classified by morphotype and

ALLAN D. CEMBELLA AND F. J. R. TAYLOR

location of origin (Table 1) are represented in Fig. 1. The zymograms reveal that a high degree of enzymatic polymorphism exists within the group. Using the similarity coefficient of Jaccard [31], Sj, with Sj = a/(a+u), where a is the number of matches and u is the number of mismatches, when isolate zymogram band patterns are compared pairwise, a similarity matrix (Table 2) was generated. Based upon the coincidence of electrophoretic bands, multiple molecular forms of all enzymes were identified. For AlaDH, at least two bands were detected (in isolates 71 and 412), with a maximum of eight bands found for isolate 180 (Fig. 1). Comparison of similarity coefficients (5"3) (Table 3) for AlaDH shows that the isolates do not group closely by geographical location or morphotype, although a slightly higher level of relatedness is expressed within the catenelloid group and the English Bay isolates, than when the zymogram data are pooled for all isolates. For GDH, 14 electrophoretically distinguishable forms were evident, with polymorphism apparent for most isolates (Fig. 1). Clonal isolates 403, 404 and 406, representing a contemporaneous population from English Bay, revealed an electrophoretically identical triple-banded pattern. Sj values for GDH indicate little apparent correlation with morphotype, but the English Bay isolates are substantially more closely related to each other than when all isolates are considered together (Table 3). The G6PDH zymograms exhibited polymorphism for most isolates, but 180, 255, 407 and 355 were monomorphic. Interestingly, the two European tamarensis isolates, the classic tamarensis from Plymouth, U.K. (183) and a smaller tamarensold form from Laguna Obidos, Portugal (253), were electrophoretically indistinguishable for this enzyme. Nevertheless, neither the entire group of tamarensoid morphotypes nor the English Bay isolates were strongly correlated for G6PDH isozymes (Table 3). Four isolates, 355, a catenelloid morphotype, and 400, 401 and 403, which are tamarensis-like, showed a monomorphic pattern for HBDH. Twenty distinguishable bands were expressed among all of the isolates. As for AlaDH and GDH, the level of relatedness is greater within the English Bay group than for all isolates (Table 3).

Patricia Bay, B.C. Canada, Aug. 1 9 7 3

Brentwood Bay, B.C. Canada, Aug. 1973 Tamar Estuary, Plymouth U.K. June. 1957 Laguna Obidos, Portugal, 1962 Lummi Island, Washington State, U.S.A. Aug. 1976 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981 English Bay, B.C. Canada, June 1981

English Bay, B.C. Canada, June 1 9 8 2

Penn Cove, Whidbey Island, Washington State, U.S.A. 1980 Friday Harbor, San Juan Island, Washington State, U.S.A. Aug. 1982 Friday Harbor, San Juan Island, Washington State, U.S.A., Aug. 1983 Whanagarei, North Island, New Zealand, Feb. 1983

Isolate

711"

180 183t

400 401 403t 4041" 4051" 4061" 407T 409 412 402

516

3551"

Absent

Absent

Absent

Absent

Present

Absent Absent Absent Present Absent Absent Absent Absent Present Absent

Absent Present

Absent Present

Present

Ventral pore

29.38 ± 2.68

32.13±2.66

33.91 + 3.33

29.34± 2.45

30.84+6.73

33.25:1:2.34 36.04+ 2.80 38.09± 3.53 33.54+ 3.13 28.42 ± 2.80 27.50+ 3.38 28.33+2.35 30.09_+2.36 28.66+ 2.75 32.91 + 2.58

24.88+ 1.75 37.09+ 1.41

37.96 ± 3.94 36.21+ 2.17

35.84± 3.40

26.71 ± 3.02

32.50±2.23

36.46 + 3.43

32.50± 2.13

29.66± 3.19

31.66+2.58 32.00± 1.88 35.13± 2.89 33.41 ± 2.88 25.91 + 2.48 26.38+ 2.34 26.26+ 2.28 27.21 ± 2.36 25.96± 2.06 33.46± 3.34

22.38± 2.09 33.34± 3.08

37.25"+ 3.05 33.84 + 1.89

32.34+ 1.96

Transapical diameter (p/n) X ± S.D.

1.10

0.99

0.93

0.90

1.04

1.05 1.13 1.08 1.00 1.10 1.04 1.08 1.11 1.10 0.98

1.11 1.11

1.02 1.07

1.11

< 2

>4

Intermediate

Catenelloid Catenelloid Catenelloid Tamarensoid

Flattened Flattened Rounded

<2

>4

>4

< 2 < 2 <~ 2 < 2 < 2 _~ 2 < 2 < 2 < 2 < 2

Tamarensoid Tamarensoid Tamarensoid Tamarensoid Tamarensoid Tamarensoid Tamarensoid Tamarensoid Tamarensoid Intermediate

Most rounded, few flattened Rounded Rounded Most rounded, few flattened Rounded Rounded Rounded Rounded Rounded Epicones and hypocones rounded or flattened Typically epicone egg- or bell-shaped hypocone tapered--some specimens have flattened epicones and hypocones Flattened

<2

<2 <2

<2

Tamarensoid Tamarensoid

Tamarensoid Tamarensoid

Tamarensoid

Morphotype

Chain formation: typical number of cells per chain*

Rounded Rounded

Epicone highly domed, hypocone rounded:~ Rounded Rounded

Shape of epicone and hypocone

* At time of original isolation. t Clonal isolate. :l:Morphological affinities with R acatenella (-- Gonyaulax acatenella [3, 33]); specimens posses a high domed epicone and prominent ventral hypothecal ridge.

508

529

435

253t 255

Location of isolate

Apical diameter (I/m) )~+ S.D.

Ratio apical: transapical diameter

TABLE 1. LOCATION OF ORIGIN AND MORPHOLOGICAL CHARACTERISTICS OF ISOLATES OF THE PROTOGONYAULAX TAMARENSlS/CATENELLA SPECIES COMPLEX

X

Z

~0

m O

=o

-I-

ALLAN D. CEMBELLAAND F. J. R. TAYLOR

314 AlaDH

71 180 183 253 255 400 401 403 404 405 408 407 409 412 402 516 355 435 529 508

m m m m m m

im~

m )

- -

__

j ~

l

l

- -

-

l , 1 - 1

1.

= i B m

-

-

-

m

m

1

ll

)

~

~

m mIj~lI

m

l

m

GDH 71 180 183253 256400401 403404 405400 407409412 402516 365435 529506

inii

m

m

m

lD223

iii ibm

lib

inn

ii

~

,i,,

m

GGPDH

71 180183 253 255 400 401 403 404 405 406407 400 412 402516 ~5§435 529 508

mm

I

m

m

m

m ,~,

m

m

m

=

m

m m

mm m

m

m

m

m m

m m m m m m m

m

m l m m

m m

m

-m

m

m mm

FIG. 1. ZYMOGRAMS OF DEHYDROGENASESEXTRACTED FROM PROTOGONYAULAXISOLATES. AlaDH, alanine dehydrogenase; GDH, glutamate dehydrogenase; G6PDH, glucose-6-phosphate dehydrogenase; HBDH, hydroxybutyrate dehydrogenase; IDH, isocitrate dehydrogenase; MDH, malate dehydrogenase; ME, malic enzyme; SucDH, succinate dehydrogenase. Staining intensity: m , intense; ~ , moderate; ~ , weak.

ELECTROPHORETIC VARIABILITY WITHIN

PROTOGONYAULAXTAMARENS/S/CATENELLA

315

HBDH 7t 180183 253 255 400 401403 404 405406 407 4n9 412 402 ~'1163S.~435 529-~=t

m

m

m

m

m

n

m

~

__

m

m m m

--

m

m m

u

m

m m

m

m

=

m

m m

N

~

m m

m

m

m

m

m

~

m

m

m

m

IDH

71 180 183 253 255 400401403404 405 4M40"/400 41Z 402 5M 355 435 S29 508

N ,m

- - m

m

m

m

m'm~m -i

n

m

m

~--U--.N ~

u

m

m

u

m ,~,

immB

_

--

m

u

,m

m

an

m

MDH 71 180 183 253 255 400 401 403 404 4Q~ 406407 409 412 402.516 355 435 529 508

_ _

--__==__---___-____-__-__--_______-_____-=-~_~

.---..__m ~ l m u

m

FIG. 1 (continued)

m m

=---

ALLAN D. CEMBELLA AND F. J. R. TAYLOR

316

ME 71 180 183 253 255 400 401403 404 40S 406 407 409 412 402 516 355 435 529 50A

m--i)m q ~

m,'~m m n

'~

~ m

i i

"~'

~'~

"~"''~

"~"

n

r~

i

n

~

a i N umR=

i I

n

n

i

SucDH

71 180 183 253 255 400 401 4Q~ 404 405 406 407 409 412 402 516 355 435 520 508

m

i

~ i

!

m

i

e

i

i

i

I

i

| i

i

i

n

H

i a

i

H

i

U

~

H

i

i

n

H n

m

FIG. 1 (continued)

For IDH, a total of 14 unique bands were produced, with monomorphic identical bands exhibited for English Bay isolates 403, 404 and 407. Two tamarensoid isolates from Vancouver Island, Canada, yielded exactly the same quadruple-banded pattern for this enzyme. In general, however, electrophoretic variants of IDH do not appear to be related geographically or according to morphotype. Although banding patterns for MDH were highly variable among the 20 isolates, with 13 unique mobility variants, this enzyme system appeared to be the most genetically conservative

of the dehydrogenases investigated. The mean Sj value pooled for all isolates was substantially higher for MDH than for any of the other enzymes. Several pairs of isolates, 400/401 (English Bay), 403/404 (English Bay) and 255 (Lummi Island, Washington)/405 (English Bay) formed electrophoretically equivalent polymorphic groups for MDH (Fig. 1)o In addition to these pairs, two isolates from the Bay of Fundy, Canada (544/545) revealed an identical quintuple-banded pattern (data not shown). The MDH patterns correlated rather highly among the tamarensoid isolates from English Bay (mean

ELECTROPHORET1C VARIABILITY WITHIN PROTOGONYAULAXTAMARENSIS/CATENELLA

317

TABLE 2. SIMILARITY COEFFICIENTS (Sj) OF ZYMOGRAM BAND PATrERNS. Sj-a/(a+u), WHERE a-NUMBER OF MATCHES AND u-NUMBER OF MISMATCHES [31 ], FOR ALL ENZYME BANDS COMPARED PAIRWISE 180 183 253 255 355 400 401 402 403 404 405 406 407 409 412 435 508 516 529

0.17 0.14 0.15 0.28 0.09 0.14 0.46 0.23 0.09 0.16 0.21 0.12 0.17 0.17 0.20 0.15 0.19 0.37 0.28

0.22 0.22 0.16 0.20 0.14 0.23 0.25 0.16 0.17 0.17 0.19 0.11 0.16 0.14 0.19 0.10 0.25 0.26

0.35 0.15 0.11 0,16 0.18 0.24 0.22 0.13 0.16 0.22 0.14 0.24 0.21 0.22 0.19 0.23 0.42

0,09 0.17 0.17 0.22 0.19 0.14 0.09 0.12 0.13 0.12 0.18 0.14 0.24 0.26 0.30 0.33

0.09 0,24 0,31 0.32 0.23 0.27 0.24 0.29 0.20 0.22 0.18 0.21 0.10 0.38 0.16

0.10 0.13 0.16 0.15 0.13 0.10 0.10 0.17 0.13 0.18 0.13 0.14 0.18 0.21

0.60 0.44 0.26 0.31 0.18 0.27 0.19 0.26 0.20 0.12 0.16 0.30 0.30

0.49 0.32 0.32 0.20 0.27 0.19 0.23 0.17 0.18 0.16 0.38 0.30

0.38 0.46 0.30 0.38 0.21 0.33 0.26 0.18 0.23 0.32 0.35

0.60 0.42 0.53 0.36 0.38 0.17 0.23 0.16 0.33 0.18

0.31 0.52 0.39 0.36 0.18 0.19 0.23 0.38 0.16

0.30 0.29 0.23 0.16 0.21 0.13 0.28 0.21

0.20 0.32 0.19 0.13 0.12 0.26 0.19

0.22 0.19 0.19 0.13 0.22 0.14

0.36 0.16 0.16 0.18 0.21

0.10 0.15 0.17 0,11

0.07 0.24 0.21

0.21 0.22

0.25

71

180

183

253

255

355

400

401

402

403

404

405

406

407

409

412

435

508

516

TABLE 3. MEAN SIMILARITY COEFFICIENTS FOR INDIVIDUAL ENZYMES BY ISOLATE AND MORPHOTYPE*

Enzymes Isolate or morphotype All isolates Catenelloid morphotype Tamarensoid morphotype English Bay isolates

AlaDH

GDH

G6PDH

HBDH

IDH

MDH

ME

SucDH

All dehydrogenases

0.18 0:25 0.18 0.24

0.23 0.25 0.22 0.31

0.18 0.25 0.12 0.19

0.13 0.14 0.14 0.21

0.14 0.09 0.17 0.13

0.44 0.22 0.41 0.78

0.24 0.06 0.25 0.47

0.11 0.08 0.07 0.14

0.22 0.17 0.21 0.33

*See Tables 1 and 2.

Sj=0.78), but less well if the distinguishing criterion was morphotype alone. All isolates exhibited polymorphism for ME, except 508 (New Zealand), with clonal isolates 403 and 407 from English Bay showing identical triplet bands. The multiple forms of ME comprised 20 mobility variants. As with MDH, isolates from English Bay appeared to be more similar for ME, than when grouped strictly by morphotype. In particular, the similarity between ME variants within the catenella-like group was extremely low (Sj=0.06), although all such isolates originated from Washington State waters. For SucDH, 16 mobility variants were identified; isolates 71 (Patricia Bay) and 516 (English Bay) expressed the same triple-banded enzyme pattern; isolates 255 (Lummi Island, Washington) and 400, 401, 402 and 407 from English Bay showed a single identical rapidly migrating

band. The level of relatedness of SucDH isozymes was very low and did not seem to be linked to geographical origin or morphotype. Comparison of the integrated Sj values for all dehydrogenases suggests that enzyme electrophoretic patterns are not tightly coupled with the provisionally assigned morphotypes. However, there appears to be a stronger relationship between tamarensoid isolates from a given geographical region (English Bay), than when electrophoretic data are pooled for all isolates conforming to this morphotype. A linkage dendrogram, a cluster analysis based upon unweighted pair-group arithmetic average (UPGMA) values of Sj (Table 3), is presented in Fig. 2. As might be expected, contemporaneous isolates from English Bay, including 403/404 and 400/401, are most closely paired. However, the English Bay isolates do not form a

318

ALLAN D. CEMBELLA AND F. J. R. TAYLOR

UPGMA

I

V 0

0.1

b403n

I

I

407 400 BC 401 402__ 255....~. WA 516

i

' 0.2

0.3

183-'--I 529~ 253~ 180..,~ 5N~

PL WA PO BC NZ

4~s.~

WA

35:~

0.4

0.5

0.6

0.7

0.8

0.9

I

1.0

Sj FIG. 2. UPGMA LINKAGE DENDROGRAM INDICATING ELECTROPHORETICAFFINITIES WITHIN THE PROTOGONYAULAX TAMARENSIS/CATENELLA SPECIES COMPLEX, CONSTRUCTED FROM Sj VALUES FROM TABLE 2. Location of origin of isolate: BC, British Columbia; WA, Washington State; PL, Plymouth, U.K.; PO, Portugal; NZ, New Zealand.

single uniform cluster. For example, isolates 71 and 255 which are from the North East Pacific coast region, but not from English Bay, cluster with the English Bay group before isolates 409/ 412 (English Bay). Isolate 529 (Friday Harbor, Washington), with a readily distinguishable catenella-form, is most closely grouped with the two European tamarensoid forms 183 and 253. The most geographically distant isolate, 508 (New Zealand) is tamarensoid in general morphology, but possesses an unusually narrow sixth precingular plate [7] which may serve to distinguish it from more typical tamarensis. The New Zealand isolate is also clustered very late in the series. Nevertheless, 355, a catenella morphotype from Washington State appears to be the most electrophoretically remote from the English Bay tamarensoid group. Discussion

The original separation of R catenella and R acatenella from R tamarensis was based upon the fact that the epithecal plate pattern, as drawn by Lebour [32] for the description of R tamarensis (as Gonyaulax tamarensis), did not conform to those of either proposed new species [33]. Furthermore, field populations assigned to R catenella formed tong chains, a characteristic not noted for R tamarensis. Since R catenella and R

acatenella were described from the San Francisco region, as Gonyaulax catenella and G. acatenella, respectively [33], while R tamarensis was identified from the Tamar estuary, Plymouth, U.K., there was also an assumption of geographical isolation. It was later independently recognized [3, 5, 6] that the epithecal plate pattern in Lebour's iconotype of tamarensis was inadvertently optically reversed. Once corrected, the thecal patterns for catenella, acatenella and tamarensis are not substantially distinguishable. In addition, co-occurrence of populations of the catenella morphotype with the tamarensis form within the same general region, in Japan and on the west coast of North America [7, 10, 34], disproves the early previous assumption that tamarensis is restricted to the Atlantic. Loeblich and Loeblich [3] considered the presence of prominant hypothecal flanges visible under SEM as diagnostic for R acatenella, although this feature was not part of the original species description [33]. Whedon and Kofoid [33] believed that R acatenella was restricted to the open sea and that tamarensis sensu Lebour was confined to estuarine waters. Nevertheless, Prakash and Taylor [35] described a toxic bloom associated with the R acateneila morphotype in the estuarine waters of Malaspina Inlet, Strait of Georgia, British Columbia.

ELECTROPHORETIC VARIABILITY WITHIN PROTOGONYAULAX TAMARENSlS/CATENELLA

319

Both field and laboratory specimens of the viewed as definitive. While these diagnostic charProtogonyaulax species complex display a level acters may be useful in discriminating of phenotypic plasticity, particularly with respect morphological species within Japanese coastal to thecal plate features, cell proportions and waters, when the criteria are rigidly applied to chain length, which tends to blur the distinction specimens from British Columbia and Massachubetween morphotypes. Working with laboratory setts (material of D. M. Anderson) morphological cultures, we have observed that the tendency 'hybrids' with a mix of characters are apparent. for R catenella-like isolates to form long chains Furthermore, it is not clear that these features do of apically/antapically flattened cells (a classic not represent a rather continuous morphological catenella trait), is frequently lost as the division spectrum in natural populations. The results of the present electrophoretic rate declines towards the onset of the stationary growth phase. The cells may become rather iso- analysis of members of this species complex diametrical compared with field specimens, for reveals the inadequacy of presently used morwhich the apical:transapical diameter ratio is phological criteria to assess genetic variability typically less than 1:1. Conversely, isolate 254 and stability within the group. Morphological from Nelson Island, British Columbia was pro- features are phenotypic expressions of the visionally classified as a strain of R tarnarensis genome, but are presumably coded for by comresembling Gonyaulax excavata sensu Balech plex interactions between several genes, and [36], except for the absence of a ventral pore, in a may be highly susceptible to environmental previous publication [34]. This isolate would now modifications. Also, even in an apparently hombe more appropriately designated as a catenella, ogeneous environment, considerable stochastic by virtue of its angularly flattened apex and anta- variability between morphotypes of individual pex, wider than long cells and its ability to form cells, which is essentially non-genetic in nature chains [7]. Morphometric measurements of con- [38], may be expressed. Isozyme electrophoresis temporaneous English Bay isolates (Table 1) of Protogonyaulax isolates represents an attempt clearly show the potential for clonal variability in to minimize such effects and to establish intracell diameters, diameter ratios (range 0.98-1.13) generic relatedness based upon characters, i.e. and shape. primary gene products, which are directly linked The presence or absence of a ventral pore to the genome. The stability of enzyme profiles may be a useful phenotypic character in that the was established by demonstrating their reprotrait is consistently maintained within a given ducibility for a given isolate during periods in strain (personal observation). However, it is of excess of a year. Thus is was possible to 'type' little use to discriminate unequivocally between isolates according to their enzyme complement tarnarensis and catenella, since, although the pore and to compare electrophoretically classified is considered to be consistently lacking in cate- strains with fresh isolates from natural popunella, it may be present or absent in tamarensis. lations. The Protogonyaulax isolate obtained by Postek It is generally assumed that the motile vegeta;imd Cox [37] from Washington State waters for tive stage in the life history of Protogonyaulax is ;heir EM study of the thecal plate patterns of cate- haploid [39, 40]. Sexual fusion of gametes nella possessed a notch (pore) on the 1' plate. results in the formation of a quadriflagellate Conceivably, a cryptic pore may be present in planozygote, which matures to yield a thickother isolates which is not visible by light micro- walled resting cyst (hypnozygote). Since excystscopy. Strains possessing a pore can not be ment produces a single flagellated cell, desigconsidered as simple geographically isolated var- nated as a planomeiocyte, which undergoes iants, since three isolates from English Bay, 404, division to form haploid daughter cells, it is not 412 and 516, display this feature, while the rest do unreasonable to assume that meiotic division not. occurs post-zygotically. The possibility that the Use of other morphological features, such as multiple isozyme expression observed in Protothe shape and pore arrangement of the APC and gonyaulax may result from gene duplication arisposterior sulcal plate, as advocated by Fukuyo [1] ing through polyploidy cannot be confirmed or to separate catenella from tarnarensis cannot be disproved without accurate chromosomal infor-

320

mation. Chromosome counts performed on R tamarensis nuclei range from 134 to 152 [41]. Since chromosomal replication is not synchronous as in eukaryotes, the considerable observed variation in chromosome number within this species is not unexpected [42]. In theory, and usually in practice, polyploidization produces increased enzymatic diversity, permitting a selective advantage due to enhanced ability to tolerate environmental extremes and exploit traditionally unfavourable niches. Nevertheless, even in the absence of polyploidy, it is reasonable to speculate that the maintenance of high levels of allelic variability within conspecific haploid dinoflagellate populations may represent a survival strategy with evolutionary significance. The haploid strategy is based upon recurrent mutation to supply genetic variety. Since unfavourable allelic variants cannot be masked as selectively neutral recessives, as is the case with diploid heterozygotes, haploid dinoflagellates may express a multiplicity of genotypes--a less conservative strategy. It is conceivable that a large number of allelic variants coded for by the genome are selectively neutral. Enzymatic function, and thus the fitness of the organism may not be substantially influenced by such ,minor substitutions detected by electrophoresis [30]. Alternatively, in a rather disequilibrated environment, selective pressures for or against alleles may be in approximate balance with recurrent mutational rates, thereby maintaining a shifting balance of genotypes without permitting monomorphic fixation [43]. Compared with the general patterns among diploid, sexually reproducing species, the Protogonyaulax species complex isolates examined in our study revealed a remarkably high degree of genetic diversity and heterogeneity, both between and within morphotypes and between morphologically similar isolates obtained simultaneously from the same location. For sexually reproducing higher plants, genetic identity values, /(Nei's statistic) reported by Gottlieb [29] for conspecific populations were typically >0.90, while values for congeneric plant species ranged from 0.28 to 0.94, with a mean of 0.67. For vertebrate populations, mean /values calculated from Ayala [44] yielded comparable estimates for local populations (0.96), subspecies (0.80) and between species and closely

ALLAN D, CEMBELLA AND F. J. R, TAYLOR

related genera (0.53). Although Gallagher [21] found that electrophoretic variability between seasonal blooms of the diploid marine diatom Skeletonema costatum exceeded that between sibling species of Drosophila and higher plants, electrophoretic studies on other diatoms, including Thalassiosira spp. [23] and Asterionella formosa [24] indicated a high degree of genetic homogeneity. For the latter two diatoms, there was little apparent genotypic heterogeneity among clonal isolates from a given geographical location, even when isolates were obtained during different seasons or years. The degree of genetic relatedness expressed by dinoflagellates of the Protogonyaulax species complex more closely approximates values obtained for sibling species (syngens) or protozoans, such as Paramecium or Tetrahymena (data recalculated from [45]). The mean similarity coefficient (Sj), determined in our study (0.22), is comparable to values reported for other dinoflageUates. Calculations based upon the data given by Schoenberg and Trench [25] for morphologically similar strains of Symbiodinium microadriaticum yielded a mean Sj value of 0.19 (range 0.00-0.72), while Hayhome and Pfiester's study on congeneric Peridinium isolates [28] gave a mean value of 0.34 (range 0.110.75). We have deliberately resisted the temptation to assign dehydrogenase isozymes to specific gene loci at this time. Instead, we have treated each bana as an independent character state expressed by the genome. Without detailed knowledge regarding the chromosome number of our isolates, the chromosomal events of sexual recombination and the possibility of posttranslational modification, it is unwise to make sophisticated genetic inferences. Nevertheless, it is strikingly obvious that in comparison with obligately sexual diploid higher plants [29] and animals [46], and with diploid algae [17, 21, 23], the dehydrogenases of the Protogonyaulax species complex are extremely polymorphic. Similar large numbers of electrophoretic variants were reported for GDH and MDH in the haploid dinoflagellate Pendinium [28]. Although is was not possible to attribute individual isozyme bands to specific intracellular compartments, the enzyme extraction procedure adopted was apparently able to solubilize

ELECTROPHORETICVARIABILITYWITHINPROTOGONYAULAXTAMARENS/S/CATENELLA both cytoplasmic and organellar constituents. Examination of cell fragments by high p o w e r phase contrast microscopy (×1000), f o l l o w i n g intensive sonication, revealed f e w intact chloroplasts of mitochondria. In most cases, use of m e m b r a n e detergents, Triton X-100, Tween-80 and deoxycholate, alone or in various combinations at 20 m M concentration, did not substantially affect band staining intensity or resolution. Also, the ability to stain for dehydrogenases, such as SucDH and IDH, w h i c h are generally considered to be mitochondrial enzymes, and ME, w h i c h occurs in the chloroplasts of higher plants, supports the case for complete organellar disruption and enzyme extraction. Selective extraction of subcellular enzyme constituents may be possible by careful fractionation. For example, w h e n MDH isozyme patterns from cells disrupted by high intensity sonication were compared w i t h those prepared by high speed mechanical homogenization, the fastest migrating bands were present o n l y from sonicated samples. The evidence from fungi, protozoa and higher animals indicates great interspecific and intraspecific diversity in the mitochondrial genetic system [47]. It is interesting to note that the dehydrogenases w h i c h are presumably mitochondrial, SucDH and IDH, also s h o w the lowest level of genotypic similarity a m o n g the Protog o n y a u l a x isolates. This may have evolutionary significance w i t h respect to the rate of change w h i c h can occur in the genes coding for such organellar proteins. Beam e t al. [27] noted an apparent correlation between electrophoretic similarity and reproductive affinities w i t h i n the C r y p t h e c o d i n i u m cohnii species complex. M o r p h o l o g i c a l l y similar breeding groups, as defined by sexual compatibility, may represent true sibling or "biological" species exhibiting reproductive isolation [48]. A t present, it is not k n o w n if the P r o t o g o n y a u l a x tarnarensis/catenella species c o m p l e x represents a binary or multiple mating system. Since not all genetic differentiation is expressed in readily identifiable phenotypic characteristics in protists [14, 23, 24, 49], morphological features m a y be inadequate to distinguish between pairs of sibling species, geographical races or ecotypes. In general, our isozyme data s u p p o r t the conclusion of H a y h o m e and Pflester [28] that enzyme electrophoresis offers a means of biochemically

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discriminating between closely related thecate dinoflagellates for w h i c h the plate patterns are essentially identical.

Experimental Culture techniques. Unialgal isolates of the Protogonyaulax tamarensis/catenella species complex retained in the NEPCC

(North East Pacific Culture Collection, University of British Columbia) were maintained in 125 ml Erlenmeyer flasks for use as stock inocula. Cultures were grown in a natural seawater medium (28%o salinity) designated NWSP-7, a nutrient enrichment modified from ASP-7 [50] as follows: Tris and Si were omitted, the trace metal:chelator ratio was adjusted by reducing the NTA concn, to 100 p.M, Mo (as NazMoO4 • 2H20) was added, to yield a final concn of 0.5 IIM, and concns of N (as NaNo3) and P (as Na2-glycerophosphate-5H~O) were increased to 1000 p.M and 100 p.M, respectively. After autoclaving, 200 p.M NaHCO3 was aseptically added to restore pH balance to 8.2. In late exponential growth phase, stock cultures were transferred using axenic technique to 2.81Fernbach flasks containing 2000 ml of autoclaved NWSP-7 medium and incubated in a growth chamber on 14:10 h light/dark cycle at 16=, with irradiance (120 p,Em-2 s-~, Biophysical quantum meter) provided by cool white fluorescent lights. Isolate iden#'ficat~bn. Twenty isolates, including nine clones, were assigned a provisional taxonomic status within the Protogonyaulax complex on the basis of morphological characteristics (as R catenella, P. tamarensis or Protogonyaulax intermediate morphotype) and designated by the NEPCCisolate number (Table 1). Cell diameter measurements (n-30) were performed using an optical micrometer under ×400 phase contrast microscopy from the ventral view. Epifluorescence microscopic observations of intact cells treated with calcofluor, which enhances resolution of the thecal plate margins, were made to confirm the presence or absence of a ventral pore. Bacterial contamina~'on. Although every effort was made to reduce bacterial contamination and subsequent growth, relatively low numbers of bacteria were noted under phasecontrast microscopy (×1000) in all cultures. Periodic counts using the epifluorescence technique typically were less than 1 × 103 bacteria per ml, and total cell volume calculations indicated that bacteria were always less than 1% at the total dinoflagellate volume. As a control, 1.01of unialgal culture was filtered through a Whatman filter under gentle vacuum to remove the dinoflagellates,while allowing the bacteria to pass into the filtrate. The filtrate was refiltered through a 0.22 Ilm GSWP Millipore membrane to retain the bacteria, which were then collected in sterile extraction buffer and electrophoresed along with dinoflagellate extracts. As a further test of possible bacterial contamination of dinoflagellate extracts, one isolate, 255, was purified by antibiotic treatment and electrophoresed in parallel with a unialgal culture. No putative bacterial bands were ever observed. Cell harvest and storage. Cells were havested at the end of exponential growth, determined by following the in vivo fluorescence curve (Turner Designs) to a maximum value. By comparing optical cell counts with in vivo fluorescence, a conversion factor was calculated and applied to adjust the culture

322 volume harvested, such that approximately equal numbers of cells ( - 2 X 10~) were obtained for each enzyme extraction. At harvest, cell densities were typically between 1.0-5.0 x 104 per ml, depending upon the isolate. Cultures were centrifuged for 5 min at 5000 g in a refrigerated centrifuge (Sorval" GSA rotor) at 4°; the loose pellets were rinsed and resuspended in a 50-fold volume of sterile-filtered washing buffer (100 mM Tris, 1 mM EDTA, ,10mM dithiothreitol, pH 7.5 at 4°), then recentrifuged (3000 g, Sorval ~ SS-34 rotor) in pre-chilled 2 dram vials. The supernatant wash buffer was aspirated away, then the cell pellets were quick frozen in dry ice and lyophilized overnight. Vials were stored in a vacuum desiccator at --40 ° until extraction. Cells preserved in this manner retained high levels of enzyme activity over a period of several months. Enzyme extraction. Cell pellets containing aproximately 2 × 106 cells were suspended in 0.35 ml of ice-cold extraction buffer (100 mM Tris, 1 mM EDTA, 10 mM dithiothreitol, 50 HM NAD ~ and 50 ~M NADP', pH 7.5 at4 °) and sonicated (Biosonik Model III) in an ice-EtOH bath at 60% of maximum output for 90 s in 10 s bursts. This procedure minimized enzyme denaturation. Microscopic examination of sonicated extracts indicated that only minute membrane fragments and virtually no intact cells or major organelles remained. After sonication, extracts were centrifuged at 27 000 g. (Sorvar SS-34 rotor) at 4 ° for 15 rain. The supernatant was applied to the gels as a crude enzyme extract. Electrophoresis. Electrophoretic separation of isozymes was performed in 1.5 mm thick 7% polyacrylamide get (PAG), using a dual-slab vertical gel apparatus (Model VS-14, Proteus Technology), with plate dimensions of 180× 140 mm. The gel buffer was 0.4 M Tris, pH 8.8 at 4°; the electrode buffer for all enzyme systems was 0.19 M glycine, 0.05 M Tris, pH 8.3 at 4 °. The electrophoresis apparatus was refrigerated externally, with additional cTooling provided by a circulating ice-water bath. Prior to the application of enzyme extracts, gels were preelectrophoresed for 50 Vh at 2 W per gel constant power. After the addition of 10% glycerol, 80 HI of enzyme extract from each isolate, equal to approximately 120 #g protein, were loaded into parallel wells. Bromophenol blue (0.01%) was added to 10% glycerol in extraction buffer and applied to the end wells to serve as the tracking dye. A reference strain, 255, of known electrophoretic characteristics was always run in the same gel position and electrophoretic mobilities normalized to a primary band from this strain, to standardize minor discrepancies between runs. After extract loading, the power was maintained at 2 W per gel for 50 Vh to allow for protein alignment within the gel. Then power was increased to 8 W per gel (400 V maximum, 75 mA maximum) until a total of 750 Vh had elapsed. Gel staining and band scoring. Gels were stained for the following enzymes, with minor modifications of standard techniques [51, 52]: alanine dehydrogenase (AlaDH, EC 1.4.1.1), glutamate dehydrogenase (GDH, EC 1.4.1.2), glucose--6-phosphate dehydrogenase (G6PDH, EC 1.1.1.49), hydroxybutyrate dehydrogenase (HBDH, EC 1.1.1.30), NADP-dependent isocitrate deydrogenase (IDH, EC 1.1.1.42), NADP-dependent malate dehydrogenase (MDH, EC 1.1.1.37), NADP-dependent malic enzyme (ME, EC 1.1.1.40) and succinate dehydrogenase (SucDH, EC 1.3.99.1). All gels were incubated in the dark at 37° for 1 h, then at room temperature until maximum resolution was achieved.

ALLAN D. CEMBELLA AND F. J. R. TAYLOR Band migration was measured from the origin with dividers and staining intensity was recorded. A photographic record of the isozyme banding patterns was made using a light table illuminated by an incandescent tungsten floodlamp (3200°K) Gels were fixed and preserved in iso-PrOH-glyceroI-HOAc (50:20:7) mixed with an equal volume of 0.2 M staining buffer.

References 1. Fukuyo, Y. (1980) Suisan KenkyuZosho33, 1 (in Japanese). 2. Loeblich, A. R., III and Loeblich, L. A. (1979) Toxic Dinoflagellate Blooms (Taylor, D. L. and Seliger, H. H., eds), p. 41. Elsevier North-Holland, New York. 3. Loeblich, L. A. and Loeblich, A. R. III (1975) Proc.lst Int. Conf. Toxic Dinoflagellate Blooms (Locicero, V. R., ed.), p. 207. Massachusetts, Science and Technology Foundation, Wakefield. 4. Steidinger, K. A. (1983) Prog. Phycol. Res. 2, 147. 5. Taylor, F. J. R. (1975) Environ. Letters& 103. 6. Taylor, F. J. R. (1979) Toxic Dinoflagellate Blooms (Taylor, D. L. and Seliger, H. H., eds), p. 47. Elsevier North-Holland, New York. 7. Taylor, F. J. R. (1984) Seafood Toxins (Ragelis, E. P., ed.), p. 77. ACS Symposium Series 262, American Chemical Society, Washington. 8. Braarud, T. (1951) Nytt Mag. Nat. Vidensk. 88, 43. 9. Alam, M. I., Hsu, C. P. and Shimizu, Y. (1979) J. Phycol. 15, 106. 10. Oshima, Y., Hayakawa, T., Hashimoto, M., Kotaki, Y. and Yasumoto, T. (1982) Bull. Jap. Soc. ScJ~ Fish. 48, 851. 11. Schmidt, R. J., Gooch, V. D., Loeblich, A. R., III and Hastings, J. W. (1978) J. Phyco/. 14, 5. 12. Schmidt, R. J. and Loeblich, A. R III (1979) J. Mar. Biol. Assoc. U.K. 59, 479. 13. Allen, S. L. and Weremiuk, S. L. (1971) Biochem. Genet. 5, 119. 14. Borden, D., Whitt, G. S. and Nanny, D. L. (1973) J. Protozoo/. 20, 693. 15. Goncalves de Lima, V. M. Q., Roitman, I. and Kilgour, V. (1979) J. Protozool. 26, 648. 16. Tait, A. (1970) Biochem. Genet. 4, 461. 17. Cheney, D. E and Babbel, G. R. (1978) Mar. Biol. 47, 251. 18. Grant, M. C. and Proctor, V. W. (1980) J. Phycol. 16, 109. 19. Thomas, D. L. and Brown, R. M. Jr. (1970) Phycologia 9, 285. 20. Thomas, D. L. and Brown, R. M., Jr. (1970) J. PhycoL 6, 292. 21. Gallagher, J. C. (1980) J. Phyco/. 16, 464. 22. Murphy, L. S. (1978) J. Phyco/. 14, 247. 23. Murphy, L. S. and Guillard, R. R. L. (1976) J. PhycoL 12, 9. 24. Soudek, D., Jr. and Robinson, G. G. C. (1983) Can. J. Bot. 61, 418. 25. Schoenberg, D. A. and Trench, R. K. (1980) Proc. R. Soc, 207, 405. 26. Watson, D. A. and Loeblich, A. R., Ill (1983) Biochem. Syst. Ecol. 11, 67. 27. Beam, C. A., Himes, M., Daggett, P.-M. and Nerad, T. A. (1982) J. Protozool. 29, 494. 28. Hayhome, B. A. and Pfiester, L. A. (1983) Am. J. Bot. 70, 1165.

ELECTROPHORETICVARIABILITYWITHIN PROTOGONYAULAXTAMARENSlS/CATENELLA 29. Gottlieb, L. D. (1981) Progress in Phytochemistry, Vol. 7 (Reinhold, L., Harborne, J. B. and Swain, T., eds), p. 1. Pergamon Press, Oxford. 30. Johnson, G. B. (1973) Ann. Rev. Ecol. Syst. 4, 93. 31. Sneath, P. H. A. and Sokal, R. R. (1973) Numerical Taxonomy, p. 1. Freeman, San Francisco. 32. Lebour, M. V. (1925) The Dinoflagellates of the Northern Seas, Mar. Biol. Assoc. U.K. (Plymouth). 33. Whedon, W. F. and Kofoid, C. A. (1936) Univ. Calif. Pub/. Zool. 41, 25. 34. Turpin, D. H., Dobell, P. E. R. and Taylor, F. J. R. (1978) J. Phyco/. 14, 235. 35. Prakash, A. and Taylor, F. J. R. (1966) J. Fish. Res. Bd Canada 23, 1265. 36. Balech, E. (1971) Pub/. Ser~. Hidrogr. Naval Argendna 654, 1. 37. Postek, M. T. and Cox, E. R.. (1976) J. Phycol. 12, 88. 38. Spudich, J. L. and Koshland, D. E., Jr. (1976) Nature, Lond. 262, 467. 39. Anderson, D. M. (1984) Seafood Toxins (Ragelis, E. P., ed.), p. 125. ACS Symposium Series 262, American Chemical Society, Washington. 40. Beam, C. A. and Himes, M. (1980) Biochemistry and Physiology of Protozoa, 2nd ed., Vol. 3 (Levandowsky, M. and Hutner, S. H. eds), p. 171. Academic Press, New York. 41. Dodge, J. D. (1963) Bot. Mar. 5, 121.

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42. Gavrila, L. (1977) Caryologia 30, 273. 43. Nevo, E. (1983) Protein Polymorphism: Adaptive and Taxonomic Significance (Oxford, G. S. and Rollinson, D., eds), p. 239. Academic Press, London. 44. Ayala, F. J. (1983) Protein Polymorphism: Adaptive and Taxonomic Significance (Oxford, G. S. and Rollinson, D., eds), p. 3. Academic Press, London. 45. Adams, J. and Allen, S. (1975) Isozymes, Genetics andEvolution, Vol. 4 (Markert, C. L., ed.), p. 867. Academic Press, New York. 46. Shaw, C. R. (1965) Science 145, 936. 47. Mahler, H. R. (1981) Origins and Evolution of Eukaryotic Intracellular Organelles (Fredrick, J. F., ed.), p. 53. New York Academy of Sciences, New York. 48. Beam, C. A. and Himes, M. (1982) J. Protozoo/. 29, 8. 49. Corliss, J. O. and Daggett, P.-M. (1983) Prodstologica 19, 307. 50. Provasoli, L. (1963) Proc. Int Seaweed Symp. Vol. 4 (DeVirville, D. and Feldman, J., eds), p. 9. Pergamon Press, Oxford. 51 Brewer, G. J. and Sing, C. F. (1970) An Introduction to Isozyme Techniques, p. 62. Academic Press, New York. 52. Harris, H. and Hopkinson, D. A. (1976) Handbook of Enzyme Electrophoresis in Human Genetics. North-Holland, Amsterdam.