A SURVEY OF STRUCTURE IN SOME FLORIDEOPHYCIDAE

by

PAUL CHARLES RUSANOWSKI B.A., San Fernando Valley State College, 1966

A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE

in the Department of Botany

We accept this thesis as conforming to the required standard

THE UNIVERSITY OF BRITISH COLUMBIA May, 1970 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study.

I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the Head of my Department or by his representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission.

Paul Rusanowski (In absentia)

Department of Botany

The University of British Columbia Vancouver 8, Canada

Date June 8, 1970 ABSTRACT

Cell wall structure was investigated in 20 different red .

Representatives from all 4- families of the order Ceramiales and one family of the order Gigartinales were investigated. Of these, 3 genera,

Polysiphonia, Pterosiphonia and Antithamnion were investigated with regards to both the cellulosic and mucilaginous portions of the cell wall. A new staining technique utilizing a combination of ruthenium red and osmium tetroxide as a postfixation was used in the latter portion of the study. The ultrastructure of pit connections was

examined in all algae.

The inner cellulosic portion of the cell wall consists of a

reticulate pattern of microfibrils which appear densely stained, In

Pterosiphonia this cellulosic portion was found to consist of 2

layers; an inner layer of microfibrils which ensheathed individual

cells and an outer layer of microfibrils which ensheathed the entire

thallus and was in contact with the mucilaginous coat. The microfibrils

in the inner layer appear nearly cross-sectioned, while those in the

outer layer appear more longitudinally oriented to the plane of

sectioning.

The outer mucilaginous coat covers the entire thallus. It

consists of 4 layers. The first or outermost layer consists of loose

bunches of microfibrils extending out from the,second layer. The

wecond layer consists of a zone of medium electron density approximately

750 A in thickness. The third layer is wholly contained within the

second layer. It is composed of a densely staining band of microfibrils extending from a similarly staining membrane-like structure. The

fourth layer is a densely stained membrane-like structure in contact with the cellulosic portion of the cell wall. An additional layer, the

D layer, is sometimes found in the cell wall. When present it is

found in the outermost portion of the cellulosic wall and obscures the

fourth layer of the mucilaginous coat. It consists of a densely

staining amorphous material.

Investigation of the pit connection showed the occurrence of

2 stages of one basic pit structure. One stage, the single disc stage- pit structure, has been found in all algae investigated. It consists

of a solid, lenticular, membrane-bound plug situated within an aperture

in the cell wall. The plug consists of a granular material surrounded by a zone of densely staining amorphous material.

The other stage, the double disc stage pit structure, is a modification of the single disc stage. It is not found in young cells near the apex of the thallus, but only in cells which have, or are

undergoing, rapid elongation and vacuolation. This pit structure has

only been observed in axial cells of the family Ceramiaceae in the

order Ceramiales. The double disc stage pit structure differs ;from

the single disc stage in that the granular material of the plug is

segregated into 2 regions or plates, one on either side of the plug.

The central region of the plug at first appears clear but later appears

to be partially occupied by a granular to fibrillar material. The

differentiation of the double disc stage pit structure from the single

disc stage has been described. These observations are thought to support and confirm the earlier work of Jungers (25). However, his observations have been extended through the use of electron microscopy in this study. It has been proposed that the terms used in this study, single disc stage- and double disc stage pit structures, replace the terms Polysiphonia and

Griffithsia pits used by Jungers. TABLE OF CONTENTS

PAGE

List of tables ...... iii

List of plates iv

List of appendicies • v

INTRODUCTION ...... 1

LITERATURE REVIEW . 2

Cell wall . .2

Pit connections • 3

MATERIALS AND METHODS 6

RESULTS ...... 9

Light microscopy ...... 10 Electron microscopy 11 Cell wall ...... 11 Pit ultrastructure • • 13 Cytoplasm • 17 DISCUSSION 20

Cell wall 20

Pit structure 22

CONCLUSIONS 29

BIBLIOGRAPHY 32

APPENDICIES . . ' 36

Appendix I i 36 Appendix II . . 38 Appendix III ...... 39 KEY TO ABBREVIATIONS . . . . 41

(ii) LIST OF TABLES

PAGE Table I. List of algae, collection and utilization data 7-8

(iii) TABLE OF PLATES

PAGE

Plates 1-2 -Light micrographs of apical region of the thallus and pits. with. both, fresh and fixed and sectioned material • .42-43

Plates 3-4 Ultrastructure of the apical cell and derivatives . .44-4-5

Plates 5-7 Ultrastructure of the single disc stage pit 46-48

Plate 8 Ultrastructure and development of the double disc stage pit ...... 49

Plates 9-12 Ultrastructure of cytoplasmic organelles and inclusions. . . • • 50-53

Plates 13-14 Ultrastructure of the cell wall .54-55

(iv) LIST OF APPENDICIES

PAGE

I. Culture formulae 36

II. Light microscope techniques and stains 38

III. Electron microscopy technique and formulae 39

Cv) 1

INTRODUCTION

One of the unique features of the Rhodophyceae is the possession of pit connections. These pit connections are absent from the sub-class Bangiophycideae. They are a prominent feature of cell morphology in the sub-class Florideophycidae to which most of the belong. Pit connections appear oval to circular in shape and are of variable size, depending on their age. In some red algae pit connections can occupy the entire wall between two cells.

Although pit connections have been the subject of numerous investigations both their structure and function remain obscure.

Recent investigations using electron microscopy have done much toward resolving the structure of pit connections, however, opinions are still divided as to their actual chemical and physical structure.

The present studies were undertaken in an attempt to determine the structure of the cell wall and pit connection in several red algae, by means of electron microscopy. Of especial importance was an investi• gation into the structure of the outermost layer of the cell wall, the so-called pectic coat, using a previously untried staining procedure.

In addition, a comparison of pit ultrastructure between different groups of algae utilizing the same preparative procedures was undertaken.

Members of all four families of the order Ceramiales and one family of the order Gigartinales were investigated. 2

LITERATURE REVIEW

CELL WALL

The cell wall in the Florideophycidae is generally considered

to be uniform in structure. It consists of two parts: an inner

cellulosic portion and an outer pectic layer (16, 19, 28). In some

cases a cuticle is also formed (28). In Porphyra, a member of the

Bangiales, the cuticle has been shown to contain a very large amount

of protein (20). In some algae, especially members of the family

Ceramiaceae, cell walls show a distinct stratification (7, 19). The

inner cellulosic portion of the cell wall consists of a pattern of

reticulate microfibrils, which are easily seen under the electron

microscope (11, 12, 4-0, 49). The microfibrils are embedded in an

amorphous matrix (26). Preston (44) has classified wall material of

algae into three groups: Group I, cellulose I being the main component with regularly oriented microfibrils; Group II, randomly oriented

microfibrils of cellulose II, and Group III, wall composed of other

microfibrils. The microfibrils have been identified as cellulose II

in Griffithsia (37), and most red algal cell walls have been placed in

Group II of Preston's classification (12, 13). The cellulosic layer

•surrounds individual cells of the thallus (49). The mucilaginous or

pectic coat, on the other hand, covers the outer surface of.the thallus

only (12, 49). Dawes et al. (12) reported this mucilaginous coat to

consist of reticulate microfibrils embedded in an amorphous matrix; while

Bisalputra et al. (2) have been able to resolve it initio four distinct 3 layers of complex nature. Surface activities at the plasmalemma have been described and related to cell wall deposition (2).

PIT CONNECTIONS

Pit connections in the red algae have received considerable attention. One of the first workers, Schmitz, described the pit as being closed by a membrane in which two plates were situated, one on either side of the membrane (52). The plates stained densely with haematoxylin, and plasmodesmata were observed passing through the membrane separating the plates. Falkenberg, working with members of the Ceramiales, agreed with Schmitz, but could not find plasmodesmata

(17). Mangenot, using Griffithsia (Ceramiales), claimed that the pit had no closing membrane and that the cytoplasm was continuous through the pit connection,(34). Miranda, using Bornetia (Ceramiales), was of the opinion that the closing membrane was present and that there were protoplasmic connecting strands (plasmodesmata) through the region (35).

Jungers was the first to propose that the confusion in this area had probably resulted from the fact that there is more than one type of pit connection in the red algae (25). He recognized two types of pit structures, the Polysiphonia-type and the Griffithsia-type (25). The

Polysiphonia-type has two densely stained plates separated by a membrane, with no plasmodesmata passing through the membrane; the

Griffithsia-type was described as a dense, biconvex^lens-shaped body, without a closing membrane. Muldorf suggested that the plates of the

Polysiphonia-type pit were actually rings which formed part of the

cell wall (36). Kylin, using Bonnemaissonia (Nemalionales), was in 4 agreement with Muldorf's results (28). These latter two interpretations are similar to the earlier work of Mangenot (34) in that cytoplasmic continuity is maintained from cell to cell by means of the pit connection.

In 1959 the first ultrastructure studies of pit connections were published by Myers et al. (38). Using osmium tetroxide fixed material they described two types of pits. One from Rhodymenia

(Rhodymeniales), was described as an open pore on the cell wall, through which the cytoplasm was continuous. They regarded it as equivalent to

Jungers' Griffithsia-type pit. The second pit structure, from

Laurencia (Ceramiales), was closed by a cell wall membrane and was considered equivalent to Jungers' Polysiphonia-type pit. Later Dawes et al. (12), using chemically cleared and macerated cell walls from members of all the orders of the Florideophycidae, also described two types of pit connections. These were termed open and closed pits.

Open pits were described as a duct or channel between two cells, as in Rhodymenia. Closed pits, as found in Laurencia, consisted of a pit closing membrane perforated by protoplasmic strands. No attempt was made to correlate- these two pit types with those of Jungers.

In contrast to earlier electron microscopy efforts, Bouck (5), working with Lomentaria (Rhodymeniales), observed a discrete, membrane- bound, biconvex plug blocking the pit aperture. The plug was shown to have a dense rim with a less dense central region. This same type of pit structure has been observed by Bischoff (4) in Thorea (Nemaliona- les), Bisalputra et al. (2) in Laurencia (Ceramiales), Hawkins (21) and Peyriere (42) in Ceramium (Ceramiales) and Ramus (46) in

Pseudogloiophlea (Nemalionales). 5

Minor variations in pit ultrastructure have also been reported.

Bouck (5) observed papillae on the plug surface. Bisalputra et al.

(2) reported a dense amorphous material extending from the edge of the plug into the middle of the cell wall. Ramus reported the deposition of material around the edge of the pit, to produce a distinct border or ring around the aperture and plug (47).

Ramus has also described the development of a plug in

Pseudogloiophlea (46). Through cell division an incomplete septum was formed between daughter cells. Nichols et a_l. (39) reported the same type of septum formation in Compsopogon. However, no plug is formed in Compsopogon and the aperture gradually disappears (39). Plug material next condenses along flattened vesicles, of possible endoplasmic reticulum origin, which become oriented longitudinally within the pit aperture. The flattened membranes disappear as deposi• tion of plug material takes place, resulting in the formation of a solid plug blocking the aperture. 6

MATERIALS AND METHODS

In the present survey, representatives of all four families of

the order Ceramiales and one family of the order Gigartinales were

studied. These algae, as well as the collection sites and specific parts of the thalli investigated, are listed in Table I.

Material was maintained either in a cold room, at 10 C+2, or

a Percival incubator with a 12 C/10 C+l day/night temperature cycle.

Day lengths were varied seasonally from 10.to 16 hours. Cool white

flourescent lights were used for illumination, with an intensity from

10-100 ft-c. Cultures were maintained either in Provasoli ES enriched

sea water (45) or Chihara marine media II (8) (app. I). A solution of 0.1% germanium dioxide was added to all culture media (31) to

eliminate contamination of cultures.

For light microscopy either fresh material or material prepared

for electron microscopy and sectioned 0.25-0.50 u in thickness was

used. Sectioned material was stained with basic fuchsin and crystal

violet (18), toluidine blue and borax, or methyl green with pyronin

(10). The staining procedure used for these stains is listed in

appendix II. Observations were made with a Nikon Model S microscope.

Photographs were taken with a Nikon EFM camera on 35 mm Kodak Panatomic

X or IIford FP 3 films.

A variety of fixations for electron microscopy was used during

this study. Those that proved to be the most satisfactory were the TABLE I

LIST OF ALGAE, COLLECTION AND UTILIZATION DATA

Name of algae Collection site;(;s) Parts Culture - X investigated field - 0

Antithamnion pacificum Rosario Beach, Wash., apical region, X, 0 (Harvey) Kylin Sooke, Brit. Col., axial cells of Coronation Island, various ages Alaska

A_. subulatum (Harvey) Rosario Beach, Wash., as in A. pacificum X, 0 J. Agardh. Sooke, Brit. Col.

9 Coronation Island, apical region, Ceramium sp. Alaska cortical cells

Platythamnjon reversum Sooke, Brit. Col. apical region, (Setchell and Gardner) young axial cells X, 0 Kylin

Pleonosporium vancouver- Rosario Beach, Wash. apical region X, 0 ianum J. Agardh.

Dasyopsis densa Smith Rosario Beach, Wash. apical region and branchlets 0

Delesseria decipiens J. Sooke, Brit. Col. young blade X Agardh.

Membranoptera multiramosa^ Rosario Beach, Wash. young blade Gardner

Polyneura latissima Rosario Beach, Wash. apical region, X, 0 (Harvey) Kylin young blade Laurencia spectabilis Rosario Beach, Wash. apical region Postels and Ruprecht

Odonthallia floccosa Rosario Beach, Wash. apical region (Esper) Falkenberg

0_. lyallii (Harvey) Rosario Beach, Wash. apical region J. Agardh.

Polysiphonia -sp. Goronation Island, apical region Alaska

Polysiphonia pacifica Sooke, Brit. Col. all parts ex• X, 0 Hollenberg var. pacifica Rosario Beach, Wash. cept the hold• fast region

P_. pacifica var. gracilis Rosario Beach, Wash. as for P. pacifica X, 0 Hollenberg var. pacifica

P_. paniculata Montagne Rosario Beach, Wash. as in P. pacifica X, 0 var. pacifica

Pterosiphonia bipinnata Rosario Beach, Wash. apical region, X, 0 (Postels and Ruprecht) pericentral cells Falkenberg var. bipinnata

Pt. gracilis Kylin Rosario Beach, Wash. apical region X, 0 pericentral cells

Agardhiella coulteri Rosario Beach, Wash. apical region

-'•The algae were identified by the author with the use of several references (15,23,24,27,4-3,51,53).

^Reference identification material for these 2 algae was lost; positive species identification was not possible with the amount of fixed and embedded material.

^Membranoptera multiramosa was collected from Rosario Beach, Washington, on March 18, 1968. It was tetrasporic at the time of the collection. This collection is believed to be a new northward extension of the occurrence of this alga, previous to this occasion, appears to be Coos Bay, Oregon (15). 9 following:

1. 2.5% glutaraldehyde in sea water (ph 6.8 and 8.0) or 0.1 M

sodium cacodylate buffer (ph 6.8); postfixed in 1.0%

osmium tetroxide in Dalton's buffer (ph 6.8) or 0.1 M

sodium cacodylate buffer (ph 6.8).

2. 2.5% glutaraldehyde or formaldehyde (41) in 0.1 M sodium

cacodylate buffer (ph 6.8); postfixed in 1.0% osmium

tetroxide plus 0.1% ruthenium red in 0.1 M sodium cacody•

late buffer made up in distilled water (ph 6.8).

Specimens were fixed for 1 hr. either at room temperature in the laboratory or at ambient field temperature. Postfixations in either case were for 1 hr. in the laboratory. An alcohol-propylene oxide dehydration series was used, and the tissues were embedded in maraglas according to Bisalputra and Weier (3). Sections were cut with glass knives, a GeFeRi or Dupont diamond knife, using either a Porter-

Blum Mt-1 or an LKB Ultrotome I microtome. Specimens were examined,, after uranyl acetate and lead citrate (48) staining, with a Hitachi

HS-7S or a Zeiss EM-9A electron microscope.

RESULTS

In this study all species of algae were surveyed with regards to pit ultrastructure. Only three genera, Polysiphonia, Pterosiphonia and Antithamnion were used in the observations by light microscopy on the cytoplasm and cell wall, and cell wall ultrastructure. 10

LIGHT MICROSCOPY

Examination of living apices of Polysiphonia, Pterosiphonia

Antithamnion species shows the apical cell to be typically dome-shaped

(figs. 1, 2). In Polysiphonia the apical cell is sometimes hidden by trichoblast filaments (fig. 2). The size of the apical derivatives increases rapidly with the distance from the apex (figs. 2, 3). Pit connections and cellular detail are not readily apparent in the apical region of the thallus.

In sectioned material the apical cell, again, appears dome- shaped (fig. 3). Derivatives appear, at first, as very narrow tiers under the apical cell (fig. 3), which undergo rapid eleongation as the distance from the apical cell increases (figs. 3, 4, 5). Lateral division of the derivatives is noticeable within the fourth derivative

(arrow, fig. 3). Vacuoles of various sizes and a nucleus with a prominent, densely staining nucleolus are observed more easily in each derivative than in the case of fresh material (figs. 3-6). The nucleoplasm is stained less intensely than the cytoplasm, and the nucleolus appears oval to round (fig. 5) with occasional vacuoles or inclusions, which may result, in some cases, in a ring-like appearance

(fig. 6).

The pit connection in both young and mature cells of Polysiphonia and Pterosiphonia appears as an aperture through the cell wall, blocked by a plug (fig. 5). This plug appears homogeneous and stains approxi• mately the same as the cytoplasm (figs. 4, 5). Pit connections are also observed in the trichoblast filaments, of Polysiphonia (fig. 4).

In the genus Antithamnion the plug of the pit connection consists of an inner light zone surrounded by a densely staining band of material

(fig. 7). With higher magnification the densely staining band of material appears as an outer dark rim, possibly due to refraction, with an inner and somewhat less dense band of material (fig. 8). The

densely staining band of material almost completely disappears in the

mid-region of the cell wall (arrow, fig. 8). The mucilaginous coat

of the cell wall has been stained very heavily in these preparations,

whereas the cellulosic wall shows variations in intensity of staining

The middle lamella region of the wall stains similarly to the mucilag

nous coat of the cell wall (figs. 4, 7). The lamellar nature of the

wall is seen in Antithamnion but not in the other algae examined

(fig. 7).

ELECTRON MICROSCOPY

CELL WALL

The appearance of the cell wall is dependent upon the type of

fixation and staining procedure employed. With glutaraldehyde or

formaldehyde fixations followed by postfixation in osmium tetroxide,

and staining with uranyl acetate and lead citrate, the wall can be

differentiated into two phases; the outer mucilaginous coat and the

inner cellulosic wall (fig. 41). The cellulosic wall appears as an

amorphous matrix either nearly devoid of microfibrils (figs. 13, 38),

or with a very loose reticulate pattern of microfibrils present (fig.

41). The mucilaginous coat appears to be composed of four layers

(fig. 42). These are an outer layer of reticulate microfibrils of 12

medium electron density, followed by a zone of densely staining material, a region of relatively low electron density, and an innermost layer, a membrane-like structure which is firmly appressed to the cellulosic portion of the cell wall. When an osmium tetroxide- ruthenium red postfixation is used in place of the previously described postfixation wall ultrastructure is revealed in much more detail. Four regions are again distinguished within the mucilaginous coat of the cell wall. These regions will be referred to as "I, II, III, and IV"

(figs. 42, 43). The outermost layer (I) consists of loose microfibrils extending out approximately 750 A from the surface of the second layer.

The second layer (II) is a zone of medium electron density, extending from the first layer down to the fourth layer, and is approximately

750 A thick (figs. 42, 43). The third layer (III) exists entirely within the second layer. It consists of a very densely staining band of microfibrils (arrows, fig. 42) which extend outward from a similarly staining membrane-like layer. The third layer is approximately 370 A thick. The fourth layer (IV) forms the innermost portion of the mucilaginous coat. This layer appears to be a membrane-like structure of approximately 60 A thickness. It is in contact with the inner

cellulosic wall (figs. 41-43). Occasionally observed is a region of amorphous material, which can be of either medium or high electron density, located directly under the fourth layer, within the outermost portion of the cellulosic wall (figs. 38, 40). This layer has been designated the D layer and, when present, completely obscures the innermost layer of the mucilaginous coat. With the use of ruthenium red-osmium tetroxide postfixation the cellulosic wall appears to consist of a very densely staining reticulate pattern of microfibrils embedded in an amorphous matrix

(figs. 42-44). The cellulosic wall can be differentiated further, in

Pterosiphonia, into two subregions. There appears to be an inner portion which ensheaths the individual cells (figs. 42, 44). This portion of the wall.is approximately 375 mu thick. There is also distinguishable an outer cellulosic layer, approximately 700 mu thick, forming a sheath around the entire thallus (figs. 42, 44). The difference between these two portions of the cellulosic wall appears to be related to the density and orientation of microfibrils in each region. The inner portion which ensheaths the. individual cells has both fewer and less compacted microfibrils than the outer portion which ensheaths the entire thallus. This is possibly related to the stages of development of the two portions of the cellulosic wall. It could also be due to the orientation of microfibrils in these two layers of the wall in which the microfibrils in layer Ll are oblique to the plane of sectioning, whereas the microfibrils in layer L2 appear to be mostly oriented longitudinal to the plane of sectioning.

Extensive surface activity of the plasmalemma has also been observed (fig. 41). Membrane-bound droplets, approximately 35 to 90 mu in diameter, can be seen at the cell surface, attached to or at some distance from the plasmalemma (arrows, fig. 41).

PIT ULTRASTRUCTURE

Only one pit structure has been found in the algae investigated so far. However, two different stages of this pit structure have been 14 observed. The most common stage observed is exemplified by Polysi• phonia and Pterosiphonia and.will be referred to as the single disc stage pit. The other stage, referred to as the double disc stage pit, is exemplified by the genus Antithamnion and is of limited occurrence.

SINGLE DISC STAGE PIT

The single disc stage pit structure has been observed in all algae examined. This pit structure basically consists of an aperture or pore through the cell wall (fig. 13). A lenticular plug is situated within this aperture (figs. 13-23). The plug exhibits a groove around its periphery which appears to firmly position it within the aperture (figs. 13-23). A membrane of approximately 100 A thickness separates the plug from the adjacent cytoplasm (figs. 13, 14, 19).

This membrane often appears loosely appressed to the surfaces of the plug facing the cytoplasm (figs. 13-20). The membrane across the face of the plug is in contact with the plasmalemma at the rim of the pit aperture (arrows, figs. 13, 14, 20, 26).

The plug consists of a thin, dense, amorphous peripheral zone of material, possibly proteinaceous, enclosing a central region of homogeneous, finely granular material (fig. 13). The inner granular matrix consists of granules approximately 100 A in diameter. The outer, dense zone varies from 240 to 620 A in thickness in the different algae examined (figs. 13-23).

MINOR VARIATIONS IN THE SINGLE DISC STAGE PIT MORPHOLOGY

Minor variations in plug morphology have been observed. In

Dasyopsis the peripheral zone of the plug was found to be up to 1000 A 15

in thickness and less dense than the inner granular matrix (fig. 17).

A dense amorphous material is sometimes associated with the groove region of the plug and extends into the middle portion of the cell wall in Odonthallia and Laurencia (fig. 16). The shape of the groove region of the plug also varies in many of the algae examined (figs. 13,

16, 19), and is considered to be due to the plane of sectioning.

However, in the pit structures of axial cells of Polysiphonia and

Antithamnion a consistent variation is apparent. The same variation is observed in pit structures between the basal cells of lateral branches and axial cells in Antithamnion. What appears to be a ridge• like extension of the inner granular matrix into the mid-region of the cell wall is present (figs. 21-23). This ridge does not occupy the entire groove region, but is offset slightly to one side.

DOUBLE. DISC STAGE PIT STRUCTURE

A major variation in the single disc stage pit structure has been observed (figs. 24-29). It occurs only in the mature elongated axial cells of Antithamnion and Platythamnion species, which are characterized by being extremely vacuolated (figs. 7, 8). This pit structure seems to occur only in the Ceramiaceae of the Ceramiales.

The plug of the single disc stage pit structure becomes modified resulting in what I have termed the double disc stage pit structure.

This pit structure consists of an aperture through the cell wall which is blocked by a biconvex, lens-shaped plug (figs. 24-29). The fine granular plug matrix, however, appears to be segregated into two approximately equal discs or plates (figs. 24-29), distinguishing it 16 from the homogeneous plug matrix of the single disc stage pit. The two discs or plates are separated by a region of low electron density, which extends across the entire diameter of the plug (figs. 25, 28, 29).

In older cells a fibrous to granular material may be found scattered within this central region (figs. 25, 29).

DEVELOPMENT OF THE DOUBLE DISC STAGE FROM THE SINGLE DISC STAGE PIT

As the plug of the single disc stage pit increases in age, the mid-region of the plug undergoes a series of changes which are chrono• logically associated with increasing age. Initially a zone of differentiation is developed in which the.inner fine granular matrix of the plug appears to become clumped and more densely stained (fig.

26). This zone is approximately 700 to 800 A wide and extends across the entire mid-region of the plug. Transparent spaces appear first near the edges of the plug (fig. 26).and continue to develop centri- petally within this zone of differentiation. This results in the plug material being segregated into two discs or plates as described above

(figs. 25, 27-29). The newly formed surfaces of the two discs bordering the central electron transparent region appear rough and uneven with protrusions of the fine granular matrix (figs. 27, 28).

Throughout these changes the outer, dense, amorphous zone remains

intact around the periphery of the plug. Finally the electron transparent region becomes, partially to completely, occupied by a

fibrous to granular matrix of approximately the same density as the

cytoplasm of the cells connected by the pit structure (figs. 24, 25, 29). 17

CYTOPLASM

CHLOROPLASTS

Proplastids have been observed only in the apical cell (figs.

9-11). Also present in the apical cell are differentiating chloroplasts which can be distinguished from proplastids by the occurrence of inner photosynthetic lamellae (figs. 10, 11). Proplastids are oval to round and range from 0.15-1.2 u. They are bounded by a double membrane envelope enclosing a heterogeneous matrix or stroma (fig. 30). Within each proplastid there is an inner double membrane closely paralleling the envelope, which is designated the outermost photosynthetic lamella by Brown and Weier (19). Both systems of membranes are approximately

125 A thick, and are separated from one another by a distance of

300-380 A (figs. 30). The outermost photosynthetic lamella occasionally shows interruptions which become especially noticeable in developing chloroplasts (fig. 31). The granular stroma has a density similar to the surrounding cytoplasm (figs. 30, 31). There are, however, regions in the stroma of low electron density which can be quite extensive

(figs. 30-33). These regions contain DNA fibrils of 25-30 A diameter, which are partially clumped together (fig. 33).

In the process of differentiation of chloroplasts the outermost photosynthetic lamella gives rise to the inner photosynthetic lamellae by invagination (arrows, fig. 33). The newly formed photosynthetic lamellae vary in length and orientation. Developing chloroplasts range from 1.8-3.3 u in length (figs. 33, 44).

In the mature chloroplast a variable number of flat, longi• tudinally parallel photosynthetic lamellae are separated by a fairly uniform distance of 740 A (figs.35, 38). Thylakoids occasionally 18

branch or fuse with one another, especially near the tips of the chloroplast. The stroma is homogeneous but for the presence of plastoglobuli (32) and occasional electron trasparent regions.

Chloroplasts have been observed in various stages of division.

Division occurs by a simple constriction of the chloroplast envelope and the outermost photosynthetic lamella (fig. 35). The thylakoids appear to separate into two during the process of division.

An anomaly to the previously described chloroplast structure has been observed in one culture of Polysiphonia pacifica. When mature

chloroplasts from axial cells were examined the chloroplasts were usually found to contain a variable number of appressed lamellae. From two to eight lamellae appear to associate and separate randomly at various points (fig. 34-). Pericentral cells of this same culture showed relatively poor preservation of chloroplast ultrastructure.

NUCLEUS

The nucleus is round to oval in shape and from 3-6 u in diameter

(figs. 9, 37-39). It possesses a prominent, centrally located, nucleolus (figs. 9, 37-39). The apical cell is uninucleate (figs. 9,11), but the derivatives are sometimes binucleate. The nuclear matrix

consists of both fine granular and coarse granular components (figs. 37,

38). The fine granular portions of the nuclear matrix, may, in part,

correspond to chromatin (fig. 37). The nuclear envelope is approximately

220 A thick, but it may exhibit extensive swelling between the inner and outer membranes of the envelope (fig. 37). Occasionally interrup• tions in the nuclear envelope, or nuclear pores, are observed. In face 19

view these pores appear to be approximately 400 A in diameter (fig. 130.

The nucleolus, a very densely staining structure, varies in size and shape (figs. 37-39). It is usually oval to round and possesses a slightly irregular margin (figs. 9, 37-39). Present within the nucleolus are one or more regions of lower electron density with a matrix similar to the nuclear matrix (figs. 9, 37-39). These regions have been variously termed vacuoles (29), nucleolar inclusions (22) and internucleonema regions (6). In some sections such a region may give the nucleolus a distinct ring-like appearance (fig. 39). These vacuole? like or low electron density regions are common in the nucleoli of plant cells (29) and were frequently observed in the nucleoli of the algae investigated in this survey.

INCLUSIONS

Crystalline structures have been observed in both Laurencia and

Polysiphonia. These structures are absent from field material collected and fixed in winter and spring. In sectioned material crystalline structures appear quite large, ranging from 0.5-1.5 u or more in diameter, and possess three of more straight sides (fig. 36). No limiting membrane has been found surrounding them. These structures appear to be composed of a very uniform, densely staining, granular material. 20

DISCUSSION

CELL WALL

Relatively little work has been done on the ultrastructure of

the cell wall of red algae using thin sectioning techniques. Thus it

is too early to generalize on red algal wall structure. However, a useless and complicated proliferation of terminology should also be

avoided. Where differences exist between previously described species

and those under consideration an attempt has been made to relate unique

and previously described wall structures to the existing terminology.

Bisalputra et al. (2) have investigated the ultrastructure of

the pectic coat of the cell wall. They were able to distinguish four

layers or zones composing the pectic coat, which were designated,

starting from the outside, the A, B, C, and D layers. These layers

are similar but not identical to layers II and III of the pectic coat

and the D layer of the cellulosic wall described in this paper. With

a new staining procedure utilizing a combination of ruthenium red and

osmium tetroxide as a postfixation it has been possible to show the

pectic coat of the cell wall in considerably more detail. The outer•

most layer, (I), consists of loose bunches of microfibrils extending

from the surface of the second layer. The second layer, (II), described

here includes the A and C layers of Bisalputra et_ al. (2). It consists

of a zone of medium electron density, approximately 750 A in thickness.

Likewise the B layer described by Bisalputra et_ al. (2) as a heavily

staining layer is similar to the third layer, (III), presented here. 21

This B layer has been shown to consist of a densely staining band of microfibrils extending from a similarly staining membrane-like structure.

The fourth layer, (IV), a densely stained membrane-like structure in contact with the cellulosic wall, is described here for the first time.

It forms the innermost layer of the pectic portion of the cell wall;

In addition to these four layers there is a band of material occasionally seen associated with the pectic coat of the cell wall. This layer has been termed the D band, and, when present, completely obscures the fourth layer of the pectic coat. The D band consists of a densely staining amorphous material, and is analogous to the D layer described by Bisalputra et al. (2).

Recently Hanic and Craigie (20) have shown that the cuticle in

Porphyra is highly proteinaceous and comprised 1/50-1/100 of the cell wall. The pectic coat described in this work from members of the

Ceramiales constitutes a similar amount of the total thickness of the cell wall. However, it cannot be determined at this time whether the cuticle isolated by Hanic and Craigie is similar to the entire pectic coat, to just one of the layers comprising it, or is not found at all in the algae used in this study.

With the use of the ruthenium red-osmium tetroxide postfixation the reticulate nature of the microfibrils in the cellulosic layer of the cell wall is readily apparent (fig. 43). In Pterosiphonia there appears to be an orientation of microfibrils in which the inner layer of microfibrils has a slightly different orientation than microfibrils in the rest of the cell wall. This can be seen in figure 44 where the inner one-third of the cellulosic wall appears to be almost a cross- sectional orientation of microfibrils whereas in the rest of the wall 22 the microfibrils appear to be more longitudinally oriented, The microfibrils in L2 also appear to be more compact than those in LI.

This orientation of microfibrils can be explained in one of two ways.

It is possible that there is a layering of wall material resulting in two layers of differently oriented microfibrils within the cellulosic wall. Such a condition has not been reported as occurring in any of the red algae thus far studied. The other possibility is that layers

LI and L2 may reflect the relative ages of these respective -layers of the cellulosic wall. The different orientation and less compact arrangement of microfibrils in LI merely reflecting the relatively recent deposition of this portion of the cellulosic wall. This latter possibility seems likely since young and actively growing apical regions were used in this study.

PIT STRUCTURE

In this survey two morphological stages of one basic pit structure were found, the single disc stage and the double disc stage pit structures. The single disc stage pit structure consists of an aperture through the cell wall which is blocked by a membrane-bound plug. The plug consists of an amorphous peripheral zone of material of variable thickness and density, enclosing a homogeneous finely granular material. This single disc stage has been observed in all

Florideophycidae examined in this survey. The structure of the single disc stage pit structure is in agreement with reports on pit ultra- structure by Bouck (5), Bischoff (4), Bisalputra et al. (2) and Ramus

(46). 23

Besides the single disc stage pit structure a second stage, the double disc stage pit structure, is present in those members of the family Ceramiaceae of the order Ceramiales examined in this survey, with the exception of Ceramium sp. in which axial cells were not examined.

The double disc.stage pit structure differentiates from the single disc stage pit structure concurrent with rapid elongation and vacuolation.

It differs from the latter stage in that material within the plug of the double disc stage pit appears to be segregated into two discs separated by a region of low electron density. The distance at which the double disc stage pit first occurs in the axial filament has not been determined. It is not present in the apical cell nor in early derivatives or cells of lateral branchlets. It is present, however, in axial cells of Antithamnion which are at least twice as long as they are wide. The differentiation of the double disc stage pit structure occurs at a later time in the pits associated with the basal cells of lateral branches. So far, the double disc stage pit is found in only

1 cell type within one family of the order Ceramiales, which may also suggest the possibility of this stage being a fixation artefact. Such an interpretation tends to be supported by the fact that in comparison to other cells, older axial cells of the Ceramiaceae could not be adequately fixed. It should be noted, however, that regardless of the fixations used and the degree of cytoplasmic preservation, pit ultra- structure remained uniform in all red algae except those in the family

Ceramiaceae. This seems to indicate, therefore, that some sort of differentiation has taken place in the central region of the plug resulting in the double disc stage pit structure. Furthermore, the 24

differentiation of the double disc stage pit occurs over a distance of several cells, and various stages of differentiation are chronologically- associated with axial cells of different ages.

There is also evidence for the occurrence of the double disc stage pit from other researchers. Peyriere (42) has observed the ultra- structure of pits in Ceramium. Though not discussing them in her report, it appears from the published micrograph (ref. 42, fig. 7) that the pit shown agrees in appearance with an early stage in the differen• tiation of the double disc stage pit structure. Hawkins (21) has recently examined the pits in Ceramium. She was able to discern that the plug was membrane-bound. The writer considers the pit structures shown by her at the Phycological Society of America meetings held in

1968 at Columbus, Ohio also correspond to early stages in the differentiation of the double disc stage pit. These reports and the

lack of other data lend support to the restriction of the double disc

stage pit to the family Ceramiaceae. Besides the restricted occurrence

of the double disc stage pit there are two important differences

between it and the single disc stage pit. The first is that of the

relative width of theppit structure in the septum of the axial cell.

In the double disc stage pit, the pit structure occupies only about

one-eighth of the end wall, whereas in comparable cells which contain

the single disc stage pit almost the entire end wall is composed of

the pit structure. In both instances the pit connection between the

apical cell and the first derivative occupies less than one-eighth of

the septum. The second difference may be due mainly to the composition

of the plug itself. The single disc stage pit plug is composed of an amorphous peripheral zone surrounding a homogeneous finely granular material. In the double disc stage pit the finely granular material is restricted to two areas along the periphery of the plug. The central region is occupied by an electron transparent region. This region could be the result of either a loss of material during fixation, that is, a fixation artefact as discussed earlier, or an actual differentiation within the plug. In the latter case this could occur in one of two ways. Either there is a splitting of the plug due to an increase in the size of the plug without an increase in the plug material, or there is a degradation of material within the central region of the plug. Since this plug structure is limited to older axial cells and there is only a limited increase in the size of the plug it seems likely that this differentiation is due to a degradation of material in the midregion of the plug.

Earlier electron microscopic studies by Myers et al. (38) and

Dawes et al. (12) described the occurrence of two different pit struc• tures in the Florideophycidae. Myers et al. (38), using Rhodymenia and Laurencia, equated the pit structures they described to the

Griffithsia and Polysiphonia types respectively of Jungers (25).

However, the writer concurs with Bouck (5), who has pointed out that the resolution of their micrographs leaves the problem unresolved. An examination of the micrographs of Myers et al. (38) shows that the pit structure described from Rhodymenia appears to be analogous to the single disc stage pit. The micrographs of the pit structure from

Laurencia are of insufficient resolution to make comparison possible.

Dawes et al. (12) surveyed the pit structures in several members of 26 the Rhodophyta and reported two types of pits which they have termed

"open" and "closed" pits. Unfortunately they used chemically cleared and macerated cell wall material which is a technique ill-suited for this problem. Several algae studied by Dawes et al. (12) (Polysiphonia,

Ceramium, Pterosiphonia) are shown in this study to contain the single disc stage pit structure. The writer considers, therefore, that the open pit-may be analogous to the single disc stage pit structure.

There is not enough evidence to substantiate the occurrence of the closed pit reported by Dawes et_ al_. (12). It is necessary, nevertheless, to repeat their survey before any definite conclusion can be made.

Ramus (46, 47) showed that the plasmalemma was continuous from one cell to the next through the pit aperture and that the pit membrane joined the plasmalemma at the rim of the aperture. The amorphous peripheral zone was divided into two parts separated by the pit membrane (46). In the present survey the plasmalemma was also seen to be continuous through the pit aperture. With respect to the pit membrane the results are slightly different. The pit membrane shows up very clearly across the face of the plug, and is not firmly attached to it. It occupies the region immediately adjacent to the outer

•surface of the peripheral amorphous zone of the plug. It is possible that both membrane associations occur, since Ramus made his observations

on a member of the Nemalionales whereas the observations in the present survey are based on members of the Ceramiales.

In a survey of the literature on pit ultrastructure it can be

seen that the single disc stage pit occurs inijthe Nemalionales (4, 46)

and Rhodymeniales (5, 38) as well as the Ceramiales and Gigartinales reported in this survey. It seems likely, therefore, that the single 27

disc stage pit is the most common pit structure in the Florideophycidae, since its presence has been established in four of the six orders in this class. It is also evident in the present study that the single disc stage pit can be structurally modified to produce another morpho•

logically distinguishable stage - the double disc stage pit structure.

At present this double disc stage pit is restricted to one family of

the order Ceramiales.

Ramus (47) has found that the cross wall region surrounding the pit can also undergo modification to produce an additional variation

in pit morphology. He has observed the deposition of material on the

septum around the periphery of the aperture to produce a distinct

ring-like swelling surrounding the pit. This type of structure might

possibly account for the description of the pit as rings by Muldorf

(36) and Kylin (28).

It is interesting to compare the results obtained in this

survey with the earlier work of Jungers (25). He proposed that two

types of pits exist in the red algae, the Polysiphonia and the

Griffithsia pits. These pit types appear to be analogous to the single

disc stage and the double disc stage pits respectively of this survey.

It appears that the Polysiphonia and Griffithsia pits can be explained

in a manner similar to that for the single disc stage and double disc

stage pits. That is, the Griffithsia pit (double disc stage) is

differentiated from the Polysiphonia type pit (single disc stage).

There are several similarities between this survey and the work of

Jungers which seem to support such an interpretation. Firstly, the

Polysiphonia pit can occupy almost the entire septum of the end wall 28

in axial cells, whereas, in algae with pit structures similar to the

Griffithsia type, pit structures occupy only a small portion of the end wall septum. The same relationship occurs between the single disc stage and the double disc stage pits. Secondly, the Polysiphonia pit was described by Jungers from Polysiphonia and Delesseria, both shown to contain the single disc stage pit in this survey. The'Griffithsia pit was described from Griffithsia and Ceramium, both members of the

Ceramiaceae. The double disc stage pit was also described in this survey from members of the Ceramiaceae. Thirdly, by comparing the drawings of the Griffithsia pit by Jungers with the electron micrographs of the double disc pit it can be seen that the description of the double disc stage pit presented here also fits the description for the

Griffithsia pit (ref. 25, figs. 15, 16). Fourthly, the single disc stage pit described in this survey and as described by others (2, 4, 5,

46) shows quite clearly that the Polysiphonia pit consists of a single homogeneous plug blocking the pit aperture.

It is proposed that the terms "single disc stage pit" which includes Jungers' Polysiphonia pit and most pit structures described in the literature, and the "double disc stage pit," which includes

Jungers! Griffithsia pit, be adopted. The use of this terminology seems more appropriate since these terms are based upon a morphological description of pit structures rather than the group of algae in which they were first described. The terminology proposed would eliminate the use of species names for structures which are of general occurrence in the Florideophycidae. These terms also eliminate the problem arising 29

from the occurrence of more than one stage of pit structure in the same alga, as occurs in Antithamnion and Platythamnion species.

CONCLUSIONS

1. The cell wall has been described in considerable detail following

the use of ruthenium red-osmium tetroxide postfixation. The

cellulosic portion of the cell wall is composed of a reticulate

arrangement of microfibrils. In Pterosiphonia this reticulate

arrangement of microfibrils can be subdivided into two regions:

an inner region where the microfibrils appear to be mostly obliquely

sectioned and an outer and much thicker region where the micro•

fibrils are mostly longitudinally oriented. The mucilaginous or

pectic coat of the cell wall appears quite complex. It has been

found to be composed of four distinctively structured layers. In

addition, a fifth layer, which is associated with the outermost

portion of the cellulosic wall, is sometimes present.

2. Two stages of one pit structure have been described from the red

algae examined in this survey, the single disc stage and the double

disc stage pit structures.

3. The single disc .stage pit occurs in all algae investigated. Most

pit structures reported in the literature can be equated to this

stage. It occurs in four of the six orders of the Florideophycidae.

An extensive investigation of all types, except the holdfast

region, in Polysiphonia showed only the single disc stage pit. The

occurrence of minor variations in the groove region of the plug have

been reported. 30

4. The double disc stage pit has been reported and described from

ultrastructure. It has been found in older axial cells of members

of the family Ceramiaceae of the order Ceramiales, It differs in

structure from the'single disc stage pit in that the matrix of the

plug of the double disc stage pit is segregated into two discs

separated by a region of low electron, density, which is in contrast

to the homogeneous matrix of the single disc stage pit. The

differentiation of this double disc stage from that of the single

disc stage pit has been described and postulated to take place by

the degradation of plug material.

5. The plug is membrane-bound. The plasmalemma passes through the

aperture of the pit. The pit membrane is loosely appressed to the

outermost face of the plug and joins with the plasmalemma at the

edge of the pit aperture.

6. The pit structures presented here substantiate Jungers' earlier

light microscope work. His original descriptions, however, have

been clarified and extended with the use of electron microscopy.

The single disc stage and the double disc stage pits used here

correspond to Jungers' Polysiphonia and Griffithsia pits respectively.

In addition, it has been proposed that the terms "single disc stage

pit" and "double disc stage pit," based on an ultrastructural

description of these structures, replace the earlier terms used by

Jungers.

7. Cytoplasmic detail in Antithamnion, Polysiphonia and Pterosiphonia

species .was found to not differ significantly from each other and

other red algae. A crystalline structure, which differs from most other reports similar structures in plants by the absence of a surrounding membrane, has been reported in Polysiphonia and Laurencia. 32

BIBLIOGRAPHY

1. Bisalputra, T. Personal communication.

2. , P. C. Rusanowski, and W. S..Walker. 1967. Surface activity, cell wall, and fine structure of pit connections in the red alga Laurencia spectabilis. J. Ultrastruct. Res. 20: 277-289,

3. , and T. E. Weier. 1963. The cell wall of Scenedesmus quadricauda. Amer. J. Bot. 50: 1011-1019.

4. Bischoff, H. W. 1965. Thorea reikei sp. nov. and related species. J. Phycol. 1: 111-117.

5. Bouck, G. B. 1962. Chromatophore development, pits, and other fine structure in the red alga, Lomentaria baileyana (Harv.) Farlow. J. Cell Biol. 12: 553-569.

6. Bresnick, E., and A. Schwartz. 1968. Functional dynamics of the Cell. Academic Press, New York. pp. 165-216.

7. Chadefaud, M. 1962. Sur quelques details de 1'organisation morphologique des parois cellulaires chez les Floridees filamentueses. Bull Soc. Bot. France. 109: 148-156.

8. Chihara, M. Personal communication.

9. Conn, H. J., M. A. Darrow and V. M. Emmel. 1960. ' Staining Procedures. Biological Stain Commission, University of Rochester Medical Center, Rochester, N. Y. , Wilkins Co. , Baltimore.

10. Dalton, A. J. 1955. A chrome-osmium fixative for electron microscopy. Anat. Rec. 121: 281.

11. Dawes, C. J., F. M. Scott and E. Bowler. 1960. Light and electron microscope study of cell walls of brown and red algae. Science, 132: 1663-1664.

12. :[ ••. , and . 1961. A light and electron microscope survey of algal cell walls. Amer. J. Bot. 48: 925-934.

13. Desikachary, T. V. 1960. Submicroscopic morphology of algae. In Proceedings of the Symposium on Algae, New Delhi, 1959. P_. Kachroo, ed. Indian Council of Agricultural Research. Job Press Ltd. Kanpur. pp. 70-77.

14. Dixon, P. S. Personal communication. 33

15. Doty, N. S. 1947. The marine algae of Oregon. Pt. II Rhodophyceae. Farlowia 3: 159-215.

16. Drew, K. M. 1951. Rhodophyta. In Manual of Phycology. G. M. Smith, ed. Ronald Press Co. New York. pp. 167-192.

17. Falkenberg, P. 1901. Die Rhodomelaceen des Golfes von Neapel und der angrezenden Meeresabschnitte. Fauna und Flora des Golfes von Neapel. Bd. 26, Berlin. (not seen, cited from ref. 28).

18. Fraser, B. Personal communication.

19. Fritsch, F. E. 1945. The Structure and of the Algae, volume II. Cambridge University Press, 939 pp.

20. Hanic, L. A., and J. S. Craigie. 1969. Studies on the algal cuticle. J. Phycol. 5: 89-102.

21. Hawkins, E. K. 1968. Cell contact in the red algae: the fine struc• ture of the pit connections in Ceramium diaphanum. (Abstr.) J. Phycol. 4(s): 6.

22. Hay, E. D. 1968. Structure and function of the nucleolus in developing cells. In Ultrastructure in Biological Systems. A. J. Dalton and F. Haguenau, eds. Academic Press. New York. 3: 474-483.

23. Hollenberg, G. J. 1942. An account of the species of Polysiphonia on the Pacific Coast of North America. I. Oligosiphonia. Amer. J. Bot. 29: 772-785.

24. . 1944. An account of the species of Polysiphonia on the Pacific Coast of North America. II. Polysiphonia. Amer. J. Bot.' 31: 474-483.

25. Jungers, V. 1933. Recherches sur les plasmodesmes chez les vegetaux. II. Les synapses des algues rouges. La Cellule 17: 1-28.

26. Kreger, D. R. 1962. Cell walls. In Physiology and Biochemistry of the Algae. R. A. Lewin, ed. Academic Press. New York, pp. 315-336.

27. Kylin, H. 1925. The marine algae in the vicinity of the biological station at Friday Harbor, Washington. Lunds Univ. Arsskr. N. F. Avd. 2, 21(9): 1-87.

28. 1956. Die Gattungen Der Rhodophyceen. CWK Gleerups Forlag. Lund. 673 pp. 34

29. Lafontaine, J. G. 1968. Structural components of the nucleus in mitotic plant cells. In Ultrastructure in Biological Systems. A. J. Dalton and F. Haguenau, eds. Academic Press. New York. 3: 152-196.

30. Leak, L. V. 1967. Fine structure of the'mucilaginous sheath of Anabaena sp. J. Ultrastruct. Res. 21; 61-74.

31. Lewin, J.. 1966. Silicon metabolism in . V. Germanium dioxide, a specific inhibitor of diatom growth. Phycologia 6: 1-12.

32. Lichtenthaler, H. K. 1968. Plastoglobuli and the fine structure of plastids. Endeavor 27: 144-149.

33. Luft, J. H. 1965. Fine structure of capillaries: the endocapillary layer. Anat. Rec. 151: 380.

34. Mangenot, G. 1924. Sur les communications protoplasmiques dans 1' appareil sporogene de quelques Floridees. Rev. Algol. 1: 376. (not seen, cited from ref. 19).

35. Miranda, F. 1930. Las Communicaciones interprotoplasmicas.en "Bornetia secundiflora" (J. Agardh.) Thuret. Ebenda, T. 30. (not seen, cited from ref. 28).

36. Muldorf, A. 1937. Das plasmatische wesen der pflanzlichen zellbrucken. Beih. Bot; CentralblattA. 56: 171-364. (not seen, cited from ref. 28).

37. Myers, A., R. D. Preston, F. R. S. and G. W. Ripley. 1956. Fine structure in the red algae. I. x-ray and electron microscope investigation of Griffithsia flosculosa. Proc. Roy. Soc. London, B/ 144: 450-459.

38. . 1959. An electron microscope investigation into the structure of the floridean pit. Ann. Bot. N. S. 23: 257-262.

39. Nichols, H. W., J. E. Ridgeway and H. C. Bold. 1966. A preliminary ultrastructural study of the fresh water red alga Compsopogon. Ann. Missouri Bot. Gard. 53: 17-27.

40. Northcote, D. H. 1963. The biology and chemistry of the cell walls of higher plants, algae, and fungi. In International Review of Cytology. G. H. Bourne and J. F. Danielli, eds. Academic Press. New York. 14: 223-259.

41. Pease, D. 1964. Histological Techniques for Electron Microscopy. Academic Press, New York.

42. Peyriere, M. 1963. Les plastes et l'amidon florideen chez quelques Rhodophycees. C. R. Acad. Sci. Paris 257: 730-732. 35

43. Phillips, R. C, and R. L. Vadas. 1967. Marine algae of Whidbey Island, Washington. J. Inst. Res., Seattle Pacific College. A(4). 82 pp.

44. Preston, R. D. 1959. Wall structure of marine algae. Proc. IX Intern. Bot. Congr. (Abstr.) 2: 310.

45. Provasoli, L. 1962. Haskins Laboratory Bulletin.

46. Ramus, J. 1969. Pit connection formation in the red alga Pseudo- gloiophlea. J. Phycol. 5: 57-63.

47. . 1969. Pit connection dimorphism in the red alga Pseudo- gloiophlea. J. Cell Biol. 41: 340-345.

48. Reynolds, E. S. 1963. The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. J. Cell Biol. 17: 208-213.

49. Roelofsen, P. A. 1959. The plant cell wall. In Encyclopedia of Plant Anatomy. W. Zimmerman and P. G. Ozenda, eds. Borntraeger, Berlin. 3(4).

50. Sabatini, D. D., K. Bensch, and R. J. Barrnett. 1963. Cytochemistry and electron microscopy. The preservation of cellular ultrastructure and enzymatic activity by aldehyde fixation. J. Cell Biol. 17: 19-58.

51. Scagel, R. F. 1957. An annotated list of the marine algae of British Columbia and Northern Washington. Nat. Museum Canada Bull. No. 150. 289 pp.

52. Schmitz, F. 1883. Untersuchungen uber die befruchtung der florideen. Sitzungsber. Akad. Wissensch. Berlin, I. (not seen, cited from ref. 38).

53. Smith, G. M. 1944. Marine Algae of the Monterey Peninsula California. Stanford University Press, Stanford, California, pp. 256-381. 36

APPENDIX I

CULTURE FORMULAE

I. Provasoli ES enrichment (4-5).

ES enrichment: Stock Amount of

solution stock/100 ml H20

NaN03 , . . 350 mg/100 ml 10 ml

. .. 50 ti 1 ml

Fe (as EDTA; 1:1 molar)1 25 ml

P II metals2 25 ml

. . 10 ug/100 ml 1 ml

. . 0.5 mg/100 ml 1 ml

. . 5 ug/100 ml 5 ml

mg/100 ml 5 ml

The ES enrichment solution is then filtered, using millipore filters number GSWP 047 00, and stored at 4- C. Use 2 ml. of ES enrichment for each 100 ml of filtered and steamed sea water.

1 Dissolve 351 mg. Fe(NH^)2.6H20 and 330 mg. of Na2EDTA in 500 ml. distilled water. One ml. of this solution equals 0.1 mg. Fe.

2 P II metals: directions for making 100 ml of metal mix.

boron - HgBOg 0;114 g.

iron - FeCl3.6H20 4.9 mg.

Manganese - MnS0^.4H20 16.4 mg.

zinc - ZnS01+.7H20 2.2 mg.

cobalt - CoSO^f^O 0.48 mg.

Na2EDTA 100 mg. 37

II. Chihara marine media II (8).

sea water 1000 ml.

minor elements 2 ml.

NaNOg 0.2 g,

NaH2P04.12H20 0.025 g.

minor elements:

distilled water • 1000 ml.

Na2EDTA . . . • 3.0 g.

FeCl3 .0.08 g.

MnCl2 0.12 g.

ZnCl2 0.015 g.

CoCl2 0.003 g.

CuCl2 0.0012 g.

Na2Mo01+.2H20 0.05 g.

H3B03 .0.6 g.

III. Preparation of germanium dioxide additive (14).

Ge02 (BDH) 1.0 g.

distilled water 1000 ml.

Shake intermittently for 3 days. Use 1-3 ml. per liter of pre•

pared media. 38

APPENDIX II

LIGHT MICROSCOPE TECHNIQUE AND STAINS

I. Staining technique:

Use \ - 3g micron sections.

1. soak in ethylene dichloride for 1 minute.

2. rinse in distilled water and then in 50% acetone.

3. heat slide until dry.

4. stain for 1-2 minutes - heat slowly during staining until steam

rises from the liquid on the slide.

5. rinse in distilled water 3 times.

6. heat slide until dry.

7. repeat steps 4-6 :if 2 stains are employed.

8. mount in immersion oil or permount.

II. Stains:

1. Basic fuchsin-crystal violet combination (18).

Make up 1% aqueous solutions of basic fuchsin and crystal violet, filter well before use. Treat sections with basic fuchsin and then crystal violet.

2. Toluidine blue and borax.

Make up a 1% aqueous solution of toluidine blue in a weak borax solution.

3. Methyl green with pyronin (9).

me'thyl green l.Og.

pyronin y or b 0.25 g.

95% alcohol 5.0 ml.

" glycerol 20 ml.

2% aqueous phenol 100 ml. 39

APPENDIX III

ELECTRON MICROSCOPY TECHNIQUE AND FORMULAE

I. Standard technique:

primary fixation . 1 hour

wash 3-6 times in buffer 1-3 hours

postfixation .1 hour

wash in a solution of \ buffer and Jg distilled water 2 times Jg hour

wash through a graded series of sea water (30%, 15%) to distilled water ...... Jg hour

alcohol dehydration series (30%, 50%,

70%, 90%, 3 changes of 100%) 2*g hours

50% alcohol-50% propylene oxide wash \ hour

3 changes of 100% propylene oxide 1 hour

infiltrate to 50% maraglas by w/v 4 hours

leave overnite in 50% maraglas (optional)

transferfto 75% maraglas by w/v 1 hour

transfer to 100% maraglas 2-3 hours change to fresh 100% maraglas and embed in capsules; cure at 65-70 C for 24- hours.

II. Preparation of primary fixatives:

1. neutralized glutaraldehyde.

Use 25 or 50% glutaraldehyde stock solution. Add enough barium carbonate to produce a 3% concentration in the above solution. Shake intermittently for 1-2 hours, then filter until clear. Check pH (6-7); if low, repeat above procedure. 40

2. formaldehyde (41).

Add paraformaldehyde to distilled water by weight-volume up to a concentration of 40%. Heat the solution to 65 C and add concentrated NaOH until the solution turns clear.

3. ruthenium red-osmium tetroxide fixative (modified from refs. 30, 33).

Add to a 1% osmium tetroxide solution in distilled water (or buffer made with distilled water) enough ruthenium red (w/v) to produce a final concentration of 0.1% ruthenium red. Let stand 15 minutes before use.

III. Buffers and embedding media:

1. sodium cacodylate I (after ref. 50).

sodium cacodylate 2.14 g.

distilled water 10 ml.

filtered sea water 90 ml.

Adjust the pH to 6.8 with HCl.

2. sodium cacodylate II (1).

sodium cacodylate 1 g,

distilled water 25 ml.

filtered sea water 25 ml.

Adjust the pH to 6.8 with HCl.

3. Dalton's buffer (after ref. 10).

Prepare a 5% w/v solution of potassium dichromate. Adjust the pH to 6.8 with 2.5N KOH, and dilute to 4% with distilled water. Mix 1 part of above solution with 1 part of filtered sea water. Use 1 part of this buffer and 1 part filtered sea water to make fixation buffer.

4. maraglas (3).

Maraglas 655 18.3 g.

Cardolite NC 513 7.2 g.

DMP 30 0.8 g.

Cure at 65-70 C for 24 hours. 41

KEY TO ABBREVIATIONS

I, II, III, IV - layers of the mucilaginous wall AC - apical cell AM - amorphous material associated with groove of pit CE - chloroplast envelope CH - chloroplast CR - central region in double disc stage pit CS - crystalline structure CW - cell wall, cellulosic CY - cytoplasm Dl, D2 - individual discs of double disc stage pit Dl - dictyosome E - amorphous D layer of material associated with the outermost portion of the cellulosic wall F - DNA fibrils G - groove of pit LI - inner portion of cellulosic wall L2 - outer portion of cellulosic wall M - plasmalemma MI - mitochondrion ML - middle lamella MW - mucilaginous coat N - nucleus NI - nucleolar inclusion NU - nucleolus OL - outermost photosynthetic lamella P - plug PM - pit membrane PP - proplastid PZ - amorphous peripheral zone of plug R - ridge in groove region of plug S - floridean starch grain T - tonoplast TC - trichoblast filament cell TS - transparent space V - vacuole ZD - zone of differentiation 42

PLATE 1

fig. 1 Antithamnion sp. showing general morphology of the apical region and dome shaped apical cell. Pit connections and cytoplasmic detail are not readily apparent. Living material. xl80. fig. 2 Polysiphonia sp. showing morphology of the apical region and the occurrence of trichoblast filaments. The apical cell is colorless and dome shaped. Living material. x40. fig. 3 P_. paniculata showing the apical region and apical cell in median longitudinal section. The amount of vacuolation observed increases with the distance from the apex of the thallus. Nuclei, with nucleoli, are readily visible. Glutaraldehyde fixation in sodium cacodylate buffer and stained with basic fuchsin-crystal violet. x780. fig. 4 P. paniculata showing older apical region and arrangement of trichoblast filament. Note pit connections between tricho• blast cells. The middle lamella is also clearly visible in the trichoblast cells. Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. Stained with methyl green-pyronin. x 750.

43

PLATE 2

fig. 5 P_. paniculata showing axial pit connections (single disc stage pit). Nuclei, with densely stained nucleoli, occupy the central regions of the cells. Glutaradehyde fixation in sodium cacodylate buffer and stained with basic fuchsin- crystal violet. "xl900. fig. 6 P_. paniculata showing nucleoli (arrows) which have a ring-like appearance due to nucleolar inclusions. Glutaraldehyde fixation in sodium cacodylate buffer and stained with tolui• dine blue with borax. x2000. fig. 7 A_. subulatum showing axial pit connection (double disc stage pit) and wall lamellae in a mature axial cell. Note the extremely large vacuole region. The plug occupies approxi• mately 1/8 of the cell wall. A lateral branch can be seen in cross section to the left of the axial filament. Double fixation in glutaraldehyde and osmium tetroxide in sodium . cacodylate buffer. Stained with basic fuchsin-crystal violet. x900. fig. 8 A_. subulatum showing detail of axial pit connection (double disc stage pit). The discs of the pit almost completely disappear in the groove region of the plug (arrow). Note that the plug occupies only a small portion of the cell wall. Fixation and staining as in fig. 7. x2600.

44

PLATE 3

fig. 9 Pterosiphonia bipinnata showing detail of apical cell. Note the large nucleus and nucleolus and the occurence of numerous proplastids and mitochondria. Double fixation in glutaralde• hyde and osmium tetroxide in Dalton's buffer. xl4700. fig. 10 Pt. gracilis showing detail of apical cell. Vacuolar region is small and limited to the apical region of the cell. Many proplastids and mitochondria are evident, as well as several dictyosomes. Double fixation in glutaraldehyde in sea water, osmium tetroxide in sodium cacodylate buffer. xl0500.

45

PLATE 4

fig. 11 Polysiphonia sp. showing three-celled lateral filament. Note arrows showing early cross-wall formation in the apical cell. Proplastids appear to be limited to the apical cell. Double fixation in glutaraldehyde in sea water, osmium tetroxide in Dalton's buffer. x4760. fig. 12 Pt. gracilis showing detail of derivatives of apical cell. Note the abundance of young chloroplasts. Mitochondria can also be seen scattered throughout the cytoplasm. Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. x5500.

46

PLATE 5

fig. 13 Pt. gracilis showing general morphology of the single disc stage pit. Mitochondria with tubular cristae are clearly evident in the cytoplasm. Note the nuclear pores (NP), shown in surface and oblique view in the nuclear membrane. Also shown is the attachment of the pit membrane to the plasmalemma•(arrow). Double fixation in glutaraldehyde in sea water, osmium tetroxide in sodium cacodylate buffer. xl7550.

47

PLATE 6

fig. 14 Laurencia spectabilis showing single disc stage pit morphology and pit membrane. Arrow indicates where contact between the pit membrane and the plasmalemma occurs. Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. x48000. fig. 15 P_. paniculata showing single disc stage pit between apical cell and the first derivative. Fixation as in fig. 14. X800000. fig. 16 Odonthallia lyalii showing single disc stage pit and associated amorphous material (AM) extending into the middle of the cell wall. Mitochondria and the tonoplast can also be seen. Double fixation in glutaraldehyde in sea water, osmium tetroxide in sodium cacodylate buffer. x24000. fig. 17 Dasyopsis densa showing single disc stage pit. Note that the peripheral zone (PZ) of the plug is less dense than the inner matrix of the plug. Double fixation in glutaraldehyde and osmium tetroxide in Dalton's buffer. x68400. fig. 18 P_. paniculata showing single disc stage pit from a trichoblast cell. The cytoplasm is restricted to a narrow band around the edge of the cell wall due the extremely large vacuole. Double fixation in glutaraldehyde and osmium tetroxide in sea water. x54100. fig. 19 L. spectabilis showing a tangential section through a single disc stage pit. Note the mature chloroplast and cross section of a mitochondrion. The pit membrane can be seen clearly along one side of the plug. Double fixation as in fig. 14. x22000.

48

PLATE 7

fig. 20 Polysiphonia sp. showing single disc stage pit and pit membrane. Pit membrane and plasmalemma contact can be seen at the arrow. Nucleus, plastids and mitochondria can be seen in the cytoplasm. Double fixation in glutaraldehyde in sea water, osmium tetroxide in Dalton's buffer. xl8000. fig. 21 P_. pacifica showing ridge-like extension of plug in pit of mature axial cell. The plug occupies almost the entire end wall of the cell in contrast to the mature plug in Antithamnion. Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. x33500. fig. 22 P_. pacifica var. gracilis showing ridge-like extension of plug in mature axial cell. Only one edge of the plug is shown, as the plug occupies almost the entire cell wall. Double fixation in formaldehyde and osmium tetroxide in sodium cacodylate buffer. x75700. fig. 23 A_. subulatum showing ridge-like extension of plug in young pit connection between axial cell and lateral branch. Compare this micrograph with fig. 29 showing a mature plug from an axial cell. Fixation as in fig. 22. x27000.

49

PLATE 8 fig. 24 Platythamnion reversum,showing double disc stage pit in axial ' cell. Central region appears to contain a fibrillar material. Pit structure occupies only about 1/8 of the cell wall. Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. x36000. fig. 25 PI. reversum showing an older axial cell than in fig. 24 with a mature double disc stage pit. Fibrillar material is present within this pit as well, but is not readily apparent in the micrograph. Note that the granular material is limited to 2 discs on either side of the plug (Dl, D2). The pit occupies approximately 1/8 of the cell wall. Fixation as in fig. 24. x53700. fig. 26 A_. subulatum showing early stage of differentiation of double disc stage pit from the single disc stage in an axial cell. Note the occurrence of only a thin layer of cytoplasm across the face of the plug. Transparent spaces (TS) can be seen at the edge of the plug and developing within the zone of differentiation (ZD). Double fixation in formaldehyde and osmium tetroxide in sodium cacodylate buffer. x24000. fig. 27 A. subulatum showing later differentiation stage of double disc ' stage pit. Transparent spaces (TS) extend nearly across the entire plug. Fixation as in fig. 26. x31300. fig. 28 A. subulatum showing a later stage in the differentiation of . the double disc stage pit than in fig. 27.- A transparent region extends across the entire midregion of the plug. Fixation as in fig. 26. x32400. fig. 29 A. subulatum showing mature double disc stage pit. Note the similarity of this pit to that shown in fig. 8. The central clear region is well differentiated from the peripheral granular discs (Dl, D2). A granular to fibrillar material partially fills the central region. The pit occupies approximately 1/8 of the cell wall. Fixation as in fig. 26. xl3500.

50

PLATE 9

fig. 30 Polysiphonia sp. showing detailed morphology of a proplastid. A dictyosome can be seen to the right of the proplastid. Double fixation in glutaraldehyde in sea.water, osmium tetroxide in Dalton's buffer. . x51400. fig. 31. Pt. gracilis showing young chloroplasts. Plastoglobuli can be seen in the matrix of one of the chloroplasts. Note also the tonoplast adjacent to the plastids. Double fixation in glutaraldehyde in sea water, osmium tetroxide in sodium cacodylate buffer. xM-6000. fig. 32 Pt. gracilis showing young chloroplast, mitochondrion, and dictyosome. The tonoplast is shown in the upper right portion of the micrograph. Fixation as in fig. 31. x53700.

51

PLATE 10

fig. 33 Pt. gracilis showing chloroplast detail and extensive electron transparent regions containing DNA fibrils.- Note the attach• ment of thylakoids to the outermost photosynthetic lamella (arrows). Also visible is the plasmalemma (upper left), mitochondria, and tonoplast (upper and lower right). Double fixation in glutaraldehyde in sea water, osmium tetroxide in sodium cacodylate buffer. x42000.

52

PLATE 11

fig. 34 P_. pacifica showing abnormal chloroplast ultrastructure. Note the association and . separation of lamellae (thylakoids) within the chloroplast. Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. x49900. fig. 35 P_. paniculata showing a dividing chloroplast. The thylakoids appear to have broken in two at the point of constriction. Daughter chloroplasts eventually pinch off from one another. Double.fixation in glutaraldehyde and osmium tetroxide in Dalton's buffer. x20300. fig. 36 P_. paniculata showing a crystalline structure in the cytoplasm. It appears to be not membrane bound. Numerous starch grains (S) are evident in the cytoplasm. Fixation as in fig. 34. x80000.

53

PLATE 12

fig. 37 P. paniculata showing detailed structure of the nucleus. Arrow indicates a region of fine granularity within the nuclear matrix (possibly chromatin). The densely staining nucleolus occupies a nearly central position. Double fixation in glutaraldehyde in sea water, osmium tetroxide in Dalton's buffer. xl7500. fig. 38 A_. subulatum showing the structure of the nucleus and cell wall. The nucleus, nucleolus with nucleolar inclusions, chloroplasts and mitochondria are visible in the cytoplasm. Note the layer of amorphous material (E) in the outermost portion of the cellulosic wall. Double fixation in glutaral• dehyde and osmium tetroxide in sodium cacodylate buffer. xl5000. fig. 39 P_. pacifica showing, nucleolus with nucleolar inclusion. Note the ring-like appearance of the nucleolus due to the nucleolar inclusion. Fixation as in fig. 38. xl8000.

54

PLATE 13

fig. 40 P_. pacifica showing details of the cell wall. The mucila• ginous portion of the cell wall is very densely stained and shows no detail. A less densely stained amorphous region (E) can be seen beneath the mucilaginous coat.. The cellulsoic portion of the cell wall appears as a reticulate arrangement of microfibrils. Double fixation in glutaraldehyde and ruthenium red-osmium tetroxide in sodium cacodylate buffer. xl5000. fig. 41 P_. paniculata showing details of the cell wall. Four layers can be distinguished within the mucilaginous wall. Compare the cellulosic portion of the cell wall, which shows few microfibrils, with the same portion of the cell wall in figs. 40, 42-44 in which ruthenium red-osmium tetroxide postfixation was employed. Note also the surface activity of the plasma• lemma (arrows). Double fixation in glutaraldehyde and osmium tetroxide in sodium cacodylate buffer. x24000. fig. 42 • Pt. bipinnata showing detail of cell wall and mucilaginous coat. Note the complexity of the mucilaginous coat as compared to that in fig. 41. Four layers are again visible, but in much greater detail. Two layers (LI, L2) can also be distinguished within the cellulosic portion of the cell wall. Double fixation in glutaraldehyde in sea water, osmium tetroxide-ruthenium red in sodium cacodylate buffer. x40000.

55

PLATE 14

fig. 43 Pt. bipinnata showing transverse section of cell wall. Note the reticulate arrangement of microfibrils in the cellulosic portion of the cell wall. The 4 layers of the mucilaginous coat are also visible. Double fixation in glutaraldehyde in sea water, ruthenium red-osmium tetroxide in sodium cacodylate buffer. x240Q0. fig. 44 Pt. bipinnata showing cell wall layers (Ll, L2) of the cellulosic portion of the cell wall in relation to the pericentral cells. Layer Ll can be seen to be continuous around each cell; while layer L2 is not present between cells and only covers the entire thallus. Fixation as in fig. 43. xl3200.