The Structure and Life History of Hormosira banksii
[Read before Wellington Branch, September 27, 1944; received by Editor, March 24, 1947; issued separately, April, 1948.]
Historical and Systematic.
The brown alga Hormosira banksii was first described under the name Fucus Banksii by Turner in his Fuci sive Plantarum Fucorum Generi a Botanicis Ascriptarum, Vol. I, 1808, where it was the first type described. This material was from. New Holland, collected by Menzies and Brown, of Captain Vancouver's expedition in 1791, and was presumably given to Sir Joseph Banks, for Turner writes: “For my specimens of this most extraordinary Fucus, I am indebted to the Right Hon. Sir Joseph Banks, by whose name I have called it…” The figure accompanying the description is of a poor but still recognisable specimen of Hormosira banksii.
In 1827 Bory described specimens of Hormosira which he called “Moniliformie de la Billardière.” May (1939) attributes three species to him—viz, Moniliformia Labillardieri, M. Banksii, and M. Sieberi, and Bory also mentioned M. dichotoma.
The name Hormosira was established for the genus by Endlicher, in 1836,† in his Genera Plantarum, and in 1842 Decaisne recorded the plant under the name Hormosira (Endl.) Banksii (Turn.) by which it is now recognised, also mentioning three other species—H. sieberi, H. triquetra, and H. nodularia. Oltmanns (1922) placed it, together with the genus Notheia, in the group Anomalae of the family Fucaceae.‡
Historical and systematic details are summarised in the following table:—
[Footnote] * Substance of thesis approved for the Degree of M.Sc with Honours, University of New Zealand, 1940.
[Footnote] † Hormosira Endl., Gen. plant. (1830) p. 10, is conserved against Moniliformia (Lamour. Dict. class., VII (1825) p. 71 Bory in Duperr. Voyage de la Coquille, Bot. (1826), p. 132). (See Intern. Rules Bot. Nomen., Jena, 1935, p. 86.)
[Footnote] ‡ Fritsch. 1945, considers these two genera a constituting a separate family, the Hormosiraceae.
|Date of Publication.||Author.||Genus.||Species.||Variety.|
|1867||Harvey in||Hormosira||Billardicri Mont.||Banksii|
|1909||Lucas||Hormosira||Banksii (Turn.) Decne.|
|"||sieberi (Bory) Decne.||varieties of|
|"||labillardieri (Bory) Mont.||H. Banksii|
|"||nodularia (Mert.) Decne.|
|"||articulata (Forsk.) Zan.|
|1927||Laing||Hormosira||Banksii (Turn.) Decaisne|
|1939||May||Hormosira||Banksii (Turn.) Decne.|
|(Lists many Synonyms for both spp.)||Hormosira?||articulata (Forsk.) Zan.*|
Harvey was the first to affirm that the different forms of Hormosira should be placed in one species, and in Phycologia Australica, 1860, he concludes his description of four varieties: “Whoever has seen and carefully studied this plant on its native coasts will…. fully agree with me in reducing to one species the several synonyms enumerated…. The differences in size and shape of the vesicles seem to result merely from local causes; either from the
[Footnote] * In answer to my enquiry, Dr. Nasr, Algologist, Marine Biological Station, Ghardaqa, Egypt, states that H. ? articulata (Forsk.), mentioned by Zanardini, is actually Hormophysa triquetra (L) Kütz., specimens of which he has seen in the British Museum.
depth of water at which the specimen was grown, or from exposure to the open sea, or shelter in enclosed harbours…. Intermediate forms between all the varieties may readily be found.”
In New Zealand, the genus Hormosira (Endlicher) has been accepted as monotypic, its single species being H. banksii (Turner) Decaisne. Local environmental conditions greatly affect the habit of the plant, giving rise to several recognisable types or varieties and many intermediate forms. This, together with the fact that collections from widely different localities were sent away from New Zealand, to be described from dried material, largely accounts for the synonomy which developed during last century.
Ecology (Plates 1–3).
The genus Hormosira (Endl.) is one of nineteen algal genera confined to Australasian waters (Laing, 1928), and is found in Australia, Tasmania, Lord Howe Island, Norfolk Island, the Kermadec Islands, Chatham Islands, and New Zealand.
It inhabits exposed rock platforms and tidepools between tidemarks, and is plentiful and fertile all the year round. Local environmental conditions chiefly determine the habit of the plant, large plants growing nearer the surface in deep, still pools, slender ones deep in tide-washed pools, and coarser plants in slightly stagnant pools near high-water mark. Harvey (1860) states, “The form Billardieri grows in deep water, Banksii in sheltered harbours, and sieberi and gracilis on exposed tidal rocks or in small rock pools.” Laing (1899) distinguishes two varieties: labillardieri, everywhere common between tidal limits, and sieberi, in tidal pools near high water mark.
Hormosira banksii may occur in a pure formation, even developing into a dense sward extending for acres, for which Hedley (1915) proposed the name “hormosiretum,” regarding it as comparable in ecological importance with the zosteretum. Oliver (1923) records this small-brown-alga subformation as most common on level rocks between tide-marks in harbours—e.g., near Purau, Lyttelton Harbour, and at Waitangi Bay, Chatham Islands, where it completely covers the rocks.
Hormosira banksii with Corallina officinalis, forms the most common association of algae in the mid-tide belt of the New Zealand coast. Both plants are light-tolerating and tolerant of some admixture of fresh water and mud, but Corallina can grow exposed to the strongest surf where Hormosira cannot exist. In rocky, open situations Hormosira occurs on projections and in crevices, and in shallow pools Corallina may cover the bottom, while Hormosira grows as a fringe round the edge. Young sporelings become established in Corallina beds and give rise to Hormosira plants on the rocky floor of the very shallow pools.
The chief characteristic of this association is the regular, diurnal, partial or complete exposure during low tide. Hormosira is enabled to withstand exposure of several hours partly because it contains internal reserves of water. Each receptacle* serves as a reservoir,
[Footnote] * The terms receptacle and connective in this paper replace internode and node of previous writers.
and is distended, hollow, and filled with saline solution. On exposure, water evaporates from the surface of the thallus, and is replenished in the tissues from this reservoir. The mucilage of the thallus makes the tissues drought-resistant, but in time the receptacles of some forms of Hormosira exhaust the reserve and may have the appearance of collapsed football bladders. This is evident in most young receptacles where mucilage is not yet present in large quantities and where the cavity is small. The plants quickly regain their turgidity when reached by the incoming tide. Slender plants of Hormosira also inhabit pools where they are never completely exposed, sometimes growing constantly at a depth of more than 4 feet below the surface.
Fronds of Hormosira which are never completely exposed may act as host to the Fucoid parasite Notheia anomala or to the endophyte Ectocarpus sp.
Material and Methods.
The main source of material has been Island Bay, Wellington, where the coastline is rich and varied in algal population, chiefly because of the wide uplifted coastal shelf which extends in places 100 yards or more out to sea, and abounds in deep and shallow rock pools.
Spring tides rise four and a-half feet at Wellington, the range being comparatively small as the tidal wave can sweep through Cook Strait with little interruption.
Collections have been made fairly regularly at all seasons of the year, the time being chosen to correspond approximately with low-tide conditions at the coast. Young plants were usually obtained from Corallina beds on the floor of shallow pools, older plants from pools or crevices and projections in more exposed parts, and other specimens for comparison were obtained from deep pools near the open sea or semi-stagnant pools near high-water mark. Whenever possible observations were made on fresh material brought, to the laboratory in collecting jars of sea-water (if covered in the field), or between damp newspaper (if exposed in the field).
Paraffin blocks were prepared containing: (i) small apical receptacles whole; (ii) very young plants (1 or 2 receptacles); (iii) older male and female receptacles, cut into small pieces containing 3 or 4 mature conceptacles; (iv) fertilized ova and sporelings (technique described below). Greatest success for blocks of the thallus and apices was obtained with Flemming's weak solution (Chamberlain, 1932). Chrome-acetic acid (Gibb, 1937) and Bouin's fluid were used for post-fertilization stages.
III. General Staining.
The mucilaginous parts of the thallus stained deeply with most reagents. The chief stains employed for fresh material were safranin, methylene blue, crystal violet and neutral red in dilute solutions (1% or less). Heidenhain's iron-alum haematoxylin with its modification by differentiation with acid 70% alcohol, followed
by light green in clove oil, or safranin in 50% alcohol, were used for paraffin sections.
IV. Special Methods for Gametes, Fertilization Stages and Sporelings.
1. Exudation of Sexual Products.–Two methods were employed for inducing the exudation of sexual products in the laboratory:—
i. At approximately the time of re-covering in the field, material which was exposed when collected was immersed in Petri dishes of seawater. After a short time exudation occurred, the ova acumulating on the floor of the dish beneath mature female receptacles, and the water surrounding a male plant becoming full of actively-swimming sperms.
ii. Material collected from the shore while still covered by the ebbing tide was kept immersed until approximately the time it would have been exposed in the field. It was then spread in large Petri dishes covered by fitting lids. After varying lengths of time, 5 minutes to 1 hour, exudation occurred, and the sexual products appeared on mature receptacles in mounds over each ostiole (orange in male plants, olive green in female plants).
2. Culturing for the Post-fertilization Stages.—Methods used were simple and proved satisfactory for obtaining stages of growth up to four or five months, but although sporelings were still apparently alive after a considerably longer time, little further growth was evident.
Fertilized ova were obtained by mixing ova and sperms in dishes of seawater, where they were either left undisturbed for 24 to 48 hours, or transferred to other containers or slides as required. The culture medium was seawater, which had been filtered and boiled for a short time, loss by evaporation being supplemented, if necessary, by the addition of distilled water. The medium was then aerated by shaking, or by bubbling air through for some time.
When the fertilized ova were firmly attached to the bottom, the containers were well shaken and rinsed to remove debris and unfertilized ova, and the prepared seawater was added to a depth of ¼ to ½ inch. Some cultures received no subsequent attention, but small quantities of nutrient solution (Gibb, 1937) or distilled water, were occasionally added in some cases, without conspicuous benefit. Some cultures were periodically aerated by passage of air bubbles. The most successful cultures were obtained when comparatively few sporelings were grown in a large quantity of water.
3. Embedding.—The method employed for obtaining sections of post-fertilization stages was adapted from Madge (1936). After fixing and thorough washing, the ova or sporelings were placed in groups on a clean slide, or on a thin film of agar on a slide. Cooled but still molten 3% agar solution was poured over them to a depth of about one tenth of an inch. When this had set, small blocks of agar were cut, each containing a group of ova or sporelings. These blocks were carried up the alcohols, etc., into paraffin and were
convenient to handle and orientate. For cutting, the paraffin was trimmed to enclose the agar block in a thin layer of wax.
4. Smears of Sperms.—The acetocarmine method (Belling, 1921, in McClung, 1937, p. 165) was used for smears of sperms from orange male exudate or from suspensions in seawater, and sperms in suspension fixed over osmic acid vapour followed by carbol-fuchsin gave good results.
Habit (Plate 1).
The name Hormosira (from ormos–a necklace, and seira—a chain), is descriptive of the general habit of the plant which has earned for it the popular names “Venus's Necklace,” “Neptune's Necklace,” and “Grapeweed”.
Description.—Frond, moniliform and branching; varying from 2 inches to 3 feet in length; differentiated into vesicated, hollow, inflated receptacles which contain internal water reserves, very variable in size and shape, spherical, oblong, ovoid, obconic, obovate, fusiform; occasionally prolonged into cylinders 2 to 3 inches long (Harvey, 1860); coriaceous; linked by short, solid connectives. Dioecious; sexual conceptacles very numerous, distributed irregularly, sunk in the peripheral tissues of the receptacles; all conceptacles of one receptacle at same stage of development; no central cavity or conceptacles in basal receptacle. Ostioles of conceptacles open on slightly raised areas which become more pronounced on drying. Growth, apical; growing point sunk in groove or depression at apex of terminal receptacle; plant held firmly to substratum by small disc-like holdfast.
Branching (Text Fig. 1).—Commonly dichotomous, from the receptacles; sometimes regularly in two alternating planes at right angles; less commonly trichotomous, irregular, from the connective, or by meristematic growth following injury to the tissues of a receptacle.
Colour.—Hormosira exhibits considerable colour variation according to the amount of light the plant receives. Parts exposed to direct sunlight may be pale and ochraceous, and shaded surfaces are olive or darker brown.
Paraphyses.—When fronds of Hormosira are immersed, tufts of delicate hairs or paraphyses project from the ostioles up to 0.6 mm. These probably keep the conceptacles free from mud, which may accumulate on the plant in considerable quantities in some habitats.
Growing Point.—There is a pyramidal group of four large, tetrahedral, apical cells situated at the base of the depression at the apex of the terminal receptacle of each branch. (Plate 2; Text Figs. 2, 3.) Considerable quantities of mucilage accumulate in this depression, which protects the growing point. These cells are readily distinguishable from surrounding tissue by their conspicuously large size, plentiful protoplasmic contents, and distinctive shape. Each cell measures approximately 50 μ. in its widest part.
In transverse section the apical cells form a somewhat circular region equally divided into four quadrants, between which lies deeply-staining mucilage which radiates out into the surrounding tissue. In longitudinal section two cells are seen side by side, together forming an inverted triangle with the base against the floor of the apical depression.
Walls formed parallel to the outer rounded faces of the apical cells, cut off daughter cells in which the sequence of division can be traced in regularly arranged tissue immediately surrounding the apical cells. No division figures were seen in the apical growing cells, but many figures were found in the undifferentiated region of cells surrounding them. Division in the apical cells is probably simultaneous. Gruber (1896–7) records the occasional occurrence of only three apical cells.
Dichotomy (Text Fig. 4).—My observations confirm the work of Gruber and establish the method of bifurcation as follows:— Radial division of two opposite apical growing-cells and of their products (A1, A3 leads to the separation of the other two large cells (A2, A1 by a band of meristematic cells. These continue active division until A2 and A4 are widely separated, and in their ultimate positions new growing points become organised and give rise to the two branches. The apical depression in the young receptacle in which bifurcation is taking place, appears broader than when no dichotomy occurs.
Central Cavity of Receptacle.—The very young apical receptacle and the basal receptacle have no central cavity. The relationship of the cavity which appears in the slightly older receptacle, to the mucilage radiating from the apical cells, has been discussed by Gruber, who states that each of the lines terminates in a separate cavity, which enlarges as the young receptacle grows, and eventually fuses with the three other cavities to form the central lumen of the receptacle. Mollet (1880) also records “a few central threads extending from the base to the top” seen on splitting open a fresh receptacle. I have been unable to confirm this in material which I have examined. Longitudinal bands of filaments project into the cavity from the inner surface of the body wall, and in the young receptacle, while the central cavity is still small, they project nearly to the centre, but I have not found a central strand present.
Differentiation.—The apical growing-cells are constantly surrounded by a region of undifferentiated, actively-dividing cells, at the borders of which differentiation is becoming evident. The uppermost cells of this region become elongated, and continue peri—and anticlinal division to form the meristoderm, the palisade-like cells of the epidermal “limiting” tissue (Bower, 1880). Beneath these, several layers of cells, becoming rounded or irregular in shape, give rise to the cortical layers of the thallus. The medullary filaments are differentiated from the lower portions of this apical region.
Mucilage.—The whole thallus of Hormosira is covered with a thin, deeply staining mucilaginous layer which, no doubt, is a factor in conserving moisture in the tissues of the plant when it has to endure regular, diurnal, low-tide exposure. Quantities of mucilage
also accumulate in the conceptacles as they approach maturity, and cells of the cortex and medulla lie embedded in a gelatinous matrix (MA.) which tends to make the tissues drought-resisting.
Cuticle (Text Figs. 7, 8).—A colourless, transparent cuticle overlies the outermost cellular layer of both receptacle and connective. It is thin on the receptacle but thick on the connective. It exhibits a great tendency to become detached, and large, delicate sheets may slough off or remain only partially adhering to the receptacles. In surface view the cuticle is divided into polygonal areas corresponding to the surface outline of the cells immediately beneath. This, together with the thickness and cellular appearance of the cuticle on the connective probably led Mollet (1880) to describe this layer as cellular. It is not cellular, but there are downward projections between the outer margins of the cells over which the cuticle lies.
Epidermal “Limiting” Tissue or Meristoderm (Text Figs. 5, 6). Several layers of small, oblong, palisade-like, polygonal cells regularly arranged in columns, constitute the chief assimilating layers of the body wall. This tissue corresponds with the “limiting tissue” of Bower (1880) and is like a many-layered, photosynthetic epidermis. It is present in both receptacle and connective, but in young tissues there is usually only one layer of oblong cells. Peri- and anticlinal divisions in these cells give rise ultimately to three or four layers. A transverse wall cuts off a basal cell (b.) and this is followed by a vertical wall in the upper daughter cell, giving rise to a characteristic arrangement of twin palisade-like cells overlying one basal cell. The basal cell does not divide further, but subsequent transverse divisions in the other cells give rise to regular columns, two of which may be subtended by one rounded basal cell.
This tissue contains many phaeoplasts lining the walls, except in the outermost layer, where the phaeoplasts surround the nucleus near the base of the cells.
Fig. 1—Types of branching, × 1–3. Fig. 2–L.S. apical growing point, showing two of the four large apical cells surrounded by undifferentiated cells. × 222. Fig. 3—T.S. apical growing point, showing mucilage radiating from between four conspicuous apical cells, × 233. Fig. 4—T.S. early stage of dichotomy × 222 (A 1 — 4, see Text); Camera lucida drawing from two consecutive sections. Fig. 5–T.S. older receptacle wall, × 265 (cuticle not shown). Fig. 6—T.S. older receptacle wall, × 148 (semidiagrammatic). Fig 7—Cuticle sloughed off receptacle, folded showing lower surface in profile, × 222. Fig. 8—Thick cuticle of connective, × 148 Fig. 9—T.S medulla of young receptacle, × 148. Fig 10—Cells of inner medullarv filaments A and B, from innermost longitudinal bands, × 33 and 83 respectively. C, occasionai secondary filament, × 83. Fig. 11–Distal half of receptacle somewhat dried to show longitudinal cobweb-like bands of medullary filaments, × 15.
aa., twin columns of “limiting” cells; b. basal cell; CU. cuticle; M, mucilage radiating from apical cells, MA. matrix: MS. mucilaginous sheath; p. pits between cells of cortex: V. vacuolate eytoplsm; C. cortex; L, “limiting” tissue meristoderm; Mu. mucilage overlying cuticle; MED, medulla.
Cortex (Text Figs. 5, 6).—Beneath the “limiting” tissue are many layers of rounded cells becoming larger and more irregular in shape towards the medulla. These cortical cells contain some phaeoplasts in the cytoplasm that lines their walls. The large cells of the inner cortical layers lie embedded in a pectic gelatinous matrix. Cells of the cortex are in communication by pits in their
walls, closed only by the middle lamella. The cortex is present in both receptacle and connective, but in the connective the cells are more regular in shape and arrangement.
Medulla (Text Figs. 5, 6, 9–11).—Internal to the cortex in the receptacle is an extensive layer of branching and anastomosing, tangled filaments, the cells of which in young receptacles are short and wide, but which become elongated in older regions. Some of the filaments end blindly in the matrix in which they lie embedded. Hypha-like filaments pass in from the innermost cortical cells and merge with the medullary filaments, and the matrix is continuous with that of the inner cortex.
The centre of the connective is occupied by a dense mass of parallel, longitudinal filaments of cylindrical cells four or five times as long as they are wide. Occasional lateral communication takes place between adjacent filaments and some branching occurs. The filaments lie in a gelatinous matrix.
At proximal and distal ends of the connective these filaments pass into the receptacles where they are grouped in longitudinal bands on the inner surface of the body-wall and project into the cavity. These filaments are more loosely connected than in the connective and are surrounded by individual mucilaginous sheaths. Some branching and lateral communication occurs, and occasional narrower secondary filaments with wide sheaths may twine amongst them. When a receptacle is cut transversely or longitudinally and allowed to dry for a few hours, the longitudinal medullary filaments appear as fine, white cobweb-like bands radiating from the insertion of the connective. (Text Fig. 11.)
These filaments of cells are more or less continuous throughout the plant, and the function of translocation is suggested by their continuous filamentous structure. The resemblance to sieve tubes has been noted (Delf, 1939) in filaments of the allied types Bifurcaria (Rees, 1933), Ascophyllum (Hick, 1885), Fucus, and Marginariella, etc., but medullary filaments of Hormosira do not show sieve plates, slime strings, or callus pads. Assimilation is carried on by all parts of the thallus and reserves are not stored in any particular region.
Holdfast.—Hypha-like filaments from the medulla of the young plant grow downward and outwards, supplementing and replacing the rhizoids of the sporeling. They give off numerous lateral branches, until a dense interweaving mat of filaments forms the small discoid holdfast. Some of the cells of these filaments have dense, orange or brown-coloured granular contents. Filaments from the holdfasts of adjacent young plants growing close together on the sub-stratum, mingle freely and may form a strong common, basal region of attachment.
Development of Cryptostomata and Conceptacles. (Text Figs. 12–17.)
The development of the conceptacles in Hormosira is largely in accordance with Bower's Theory (1880) based on Fucus spp. This conclusion is supported by observations of Getman (1914).
Youngest stages of the cryptostomata or “hairpits” are found
in the small apical receptacle. A cell in the superficial layer, quite near the apical growing-cells, often within the apical depression, temporarily ceases active division, and because of continued growth of the surrounding tissue, becomes sunk slightly below the surface. It becomes flask-shaped and its neck is surrounded by mucilage. The mouth of the tiny pit becomes filled with a “tongue” of mucilage which as adjacent cells contribute, accumulates in considerable quantities in the enlarging depression. The initial cell (i) divides by a transverse wall, cutting off a basal cell (b.), radial divisions of which give rise to cells which form the greater part of the floor and walls of the cryptostoma, while the cells nearer the mouth of the depression are from the superficial “limiting” layer. While the depression is still comparatively shallow, hairs arise on its floor. They increase in length not only by the multiplication of large basal cells, but also by great elongation of all the cells, until the hairs project a considerable distance through the ostiole. The cavity of the cryptostoma is all the time enlarging by continued multiplication of the cells of its floor and walls, and, because of the overgrowth of the superficial cells of the thallus, the ostiole and neck remain narrow. The cryptostoma becomes a narrow flask-shaped pit, from
Conceptacle Development—(Text Figs. 12–19).
Figs. 12–16—Stages in development of conceptacle from superficial initial cell, × 222. Fig. 17—Floor of transition stage between cryptostoma and conceptacle, × 338. Figs. 18 and 19—Exudation of antheridia and ova from ostioles, × 37.
the floor of which grow numerous unbranched, multicellular hairs which fill the cavity and project in a delicate tuft through the ostiole.
At this stage the cryptostomata, as such, may be called mature. They are homologous with cryptostomata on vegetative parts of many other Fucaceae, but in Hormosira, all the cryptostomata are potential sexual conceptacles, and they now enter a transitional stage.
The cavity enlarges and new papillae arise from the floor and walls among the original hairs. These give rise to hairs of a different kind, the paranemata, and gradually the original ones are lost. There is still no means of distinguishing between male and female conceptacles.
If the conceptacle is to be male, segmentation of the paranemata may continue until elaborately branched structures bearing numerous antheridia are formed.
If the conceptacle is to be female, the paranemata may become branched or unbranched and other papillae arise amongst them and give rise to the oogonia.
Thus, the cryptostoma, which is a juvenile stage of the conceptacle, originates from a slightly modified superficial cell of the thallus close to the apical growing-cells, and the bulk of the floor and sides of the cryptostoma is formed from the “basal cell,” with some contribution from the “limiting” layer.
Mature Female Conceptacle.
The mature female conceptacle is flask-shaped, opening to the exterior by a narrow neck and a pore, the ostiole. It is sunk in the wall of the receptacle to a depth of 0 8–0·9 mm. Through the ostiole project long, delicate, multicellular, unbranched hairs or paraphyses, similar to those projecting from the cryptostomatal stage in younger receptacles, but these ar [ unclear: ] se, not from the floor of the conceptacle, but from a restricted area of the wall immediately at the base of the neck. They exhibit basal growth and may project 0 6 mm. beyond the ostiole.
Entirely different hairs, the paranemata, arise from all parts of the walls of the conceptacle, even between the bases of the paraphyses. These paranemata are delicate, almost colourless, branched or unbranched, and multicellular. Branches originate as peg-like outgrowths from the anterior ends of the cells, and they may grow quite long before being cut off by a wall. The paranemata grow in tufts and may project more than 0.4 mm. into the conceptacle, the cavity of which becomes largely filled with them.
In a preparation stained with dilute aqueous safranin, each paranema is found to be surrounded by a wide mucilaginous sheath associated with which are additional layers of less deeply staining mucilage (Text Fig. 25). This preparation serves to indicate the very great quantity of mucilaginous material that is present in the mature conceptacle.
The female sex organs are oogonia (Text Figs, 20–22). They arise in great numbers from the walls and floors of the conceptacle, between the bases of the paranemata. These papillae differ from papillae which will give rise to vegetative paranemata, in having
denser cell contents. A papilla divides transversely forming a stalk cell (which is never very large or conspicuous) and the oogonium mother-cell, which is at first a rounded, uninucleate cell with dense, vacuolate cytoplasm and some phaeoplasts. The oogonium mother-cell enlarges considerably, before its nucleus divides three times. The first two divisions are probably meiotic. According to Getman (1914) evanescent wall formation often takes place at the eightnucleate stage; horizontal and/or vertical walls appear, or rarely, nuclei and plastids are blocked out by walls. No spindles were seen to account for the wall formation, but Getman suggests that walls may have been formed following cleavage, as happens among certain algae and fungi. He goes on to say that it seems certain that eight eggs begin to develop and that the four-egg condition is reached by breaking down of immature eggs rather than four free nuclei. He interprets this as less removed from the Fucus condition than when four free nuclei break down. My observations of the mature oogonium confirm the fact that only four ova mature. I have frequently seen, within the exochite and released with the mature ova,
Oogonial Development—(Text Figs. 20–25).
Fig. 20—Stages in development of oogonia, × 222. Fig. 21—Typical mature oogonium, × 200. Fig. 22–Unusual oogonium containing two enlarged ova (102 μ long), and five visible undeveloped ova, × 200. Fig. 23A and B, Release of ova from exochite, × 200. Fig. 24—Release of ovum from endochite (stained methylene blue), × 3.50. Fig. 25—Short female paranema enveloped in mucilage (stained safranin), × 222.
three or four bodies measuring up to 11 μ which probably are undeveloped ova.
There is considerable growth in size while the four oospheres are being organised. The mature oogonium measures about 160 μ by 110 μ It contains four uninuclcate ova, each enclosed in an envelope. They are not perfectly spherical, the shape being conditioned by enclosure within the exochite, which is the thin, firm, pectic bounding capsule of the oogonium. Within the exochite is a small quantity of mucilage—the mesogelatin.
All stages of oogonium development are found in a single mature conceptacle at any season of the year. I have found fertilization and subsequent development more reliable in the warmer months, but it is possible to obtain stages all the year round.
Release of the Ova (Text Figs. 23, 24).—The release of ova in nature occurs soon after the plants have been covered by the incoming tide, twice every 24 hours, and laboratory experiments confirm this conclusion. On no occasion have I seen any trace of exudate on plants in the field. During exposure, great evaporation takes place, the cells of the thallus lose turgor, and a certain amount of shrinkage occurs. When the tide again covers the plant, these tissues immediately absorb water and regain turgidity. As already noted, the mature conceptacle contains large quantities of mucilage within the oogonium and enveloping the paranemata, etc. The swelling of this mucilage is instrumental in releasing the sexual products from plants which are subject to low-tide exposure.
The mesogelatin swells and eventually ruptures the exochite at the apex, releasing the four ova into the cavity of the conceptacle. The exochite usually remains attached to the stalk cell. Great numbers of ova are released in this way, some still enveloped in small quantities of mesogelatin, and each enclosed in a two-layered mucilaginous envelope. The pressure caused by the great swelling in volume of the mucilage as it imbibes water forces the ova through the ostiole, one after another. As each ovum is squeezed out it becomes elongated, but it quickly ressumes its shape. When exudation begins, the paraphyses are somewhat flattened over the thallus, but they are drawn together and then jerked apart as each group of 4, 8, or 12 ova passes out, drawing together slightly after each exudation. The products of each oogonium (or several oogonia) tend to remain together during and after their passage through the ostiole, and they may remain entangled in the paraphyses for a short time if exudation is not very active. (Text Fig. 19.)
Hormosira does not exhibit the succession of oogonial walls shown by Fucus (Bower, 1880), from which the ova are simultaneously released, naked, into the water. When released from the exochite, the four ova are enclosed in individual, two-layered envelopes, and are temporarily held together by a small quantity of mesogelatin. The layers of the enclosing envelope can be demonstrated by differential staining. When ova are observed in a medium of saltwater containing a drop of 1% aqueous solution of methylene blue, the outer layer of the envelope at first stains violet and the inner layer stains deep violet. When the ovum is released, it spins
free, leaving a path of mucilage grading from deep blue to violet, and the wall of the envelope stains purple. This differential staining is transitory, and soon the whole preparation is blue. The envelope appears to consist of an outer membrane, within which is a considerable quantity of mucilage, and I interpret them to be endochite and mucilaginous endogelatin. I consider the ovum within the oogonium to be surrounded by four layers, but only two walls: (1) the exochite, common to the four ova, within which is mucilaginous mesogelatin; (2) the individual endochite enclosing a considerable quantity of mucilaginous endogelatin.
Immediately the ovum comes in contact with the water outside the ostiole, its envelope begins to swell, and after a short time, varying from 10 seconds to several minutes, the ovum protrudes against the envelope wall, eventually rupturing it and the protoplast slips through the break. It immediately assumes a spherical form measuring 64–74 in diameter. Frequently, the final release of the ovum from its envelope is sudden enough to send it spinning as it sinks to the bottom of the containing vessel. Sometimes the ova retain their envelopes for several minutes until they may swell to 112 μ wide and 126 μ long. When great numbers of ova are exuded, many may fall to the bottom of the dish without losing their envelopes. Often, it is lost in a non-active way, by slow separation as if dissolving at one point. Exudation from one receptacle is usually completed after about 5–10 minutes.
If, in the laboratory, mounds of exuded ova were allowed to remain for some hours before immersion, little activity was shown, the ova often failed to lose their envelopes, and fertilization seldom occurred. Liberation into seawater a short time after exudation is essential to further development, as it will be seen that fertilization can only occur when the sperms have access to the free surface of the ovum.
The mechanism of release of sexual products from plants which are constantly immersed in deep pools is obscure, but all the plants I have examined have been normally fertile.
The Unfertilised Ovum (Text Figs. 40, 46).—The unfertilised ovum is a naked protoplast surrounded only by the thin cytoplasmic membrane. It is spherical, with an easily distinguishable, central nucleus, the position of which is indicated by a comparatively clear region about 12 μ in diameter, surrounded by denser and more deeply-coloured cell contents in which are many phaeoplasts and granules of reserves, etc. The ova are furnished with the means of assimilation and abundant reserves from the beginning of their independent existence, and they remain fertile for several days (up to 1 week) in a dish of seawater.
There is an outer layer of delicately alveolar cytoplasm containing few phaeoplasts, beneath which is an extensive layer of “foamy” (Farmer and Williams, 1896), vacuolate cytoplasm with many phaeoplasts lining the walls of the vacuoles. The central large nucleus, which usually has one large nucleolus, rarely another small one, lies in a dense, more “granular,” finely alveolar layer of cytoplasm containing no phaeoplasts.
Mature Male Conceptacle.
In size, shape, position in the receptacle, origin, and vegetative anatomy, the male conceptacle exactly resembles the female conceptacle.
The male sex organs (Text Figs. 26–35) are antheridia borne in great numbers on tufts of much-branched, delicate paranemata which grow from all parts of the wall, and almost completely fill the cavity of the conceptacle. The antheridia arise in place of branches from the paranemata, and from the beginning, are distinguishable from purely vegetative branches by their abundant cytoplasmic contents. Some vegetative branches grow very long and resemble the paranemata of the female conceptacle. There are similar indications of the presence of large quantities of mucilage in the male conceptacle.
The antheridium is comparatively large before its large nucleus undergoes division, and subsequent growth is in length. Six divisions occur, the first two of which are meiotic, indicated by comparison of chromosome counts from the first division in the antheridium, sperm nuclei, and dividing vegetative nuclei. The nuclei of later stages are smaller, and various stages of their division have been observed (in paraffin material) some of which are figured (Text Figs. 26–33). Division within the antheridium is simultaneous.
The antheridia are almost colourless until differentiation of the antherozoids or sperms begins, after the formation of the 64 nuclei. Then the carotin in the developing stigmata gives them a characteristic orange colour.
The mature autheridium contains 64 sperms. It has two walls, the exochite which remains attached to the paranema within the conceptacle and the endochite within which the sperms are carried through the ostiole. As in female plants, release occurs when swelling of mucilage takes place after re-covering by the flow-tide. The freed antheridia fill the conceptacle and are ejected through the ostiole, accompanied by jerking of the paraphyses at each period of active exudation. (Text Fig. 18.)
The antheridium measures up to 42 μ long and 17 μ wide. Within the endochite is a small quantity of endogelatin which imbibes water and swells. The endochite ruptures at the apex, and through this opening the sperms bulge into a globular mass up to 20 μ in diameter, enveloped in mucilaginous endogelatin. By this time the sperms are showing some activity. Sometimes the globular mass contains wriggling sperms, and usually a few sperms still within the endochite are very active. After a short time sperms near the edge of the globular mass become more active and struggle to free themselves from the mucilage, and one by one they dart away. Quite often the most active sperms inside the endochite fail to escape, probably because the remains of the endogelatin blocks the opening.
The Structure of the Sperm (Text Figs. 37–39).—The male gamete of Hormosira is somewhat pear-shaped, with one side flattened. It measures 5 or 6 μ in length and 2 μ in its widest part. It has a large nucleus surrounded by a thin layer of cytoplasm in
Antheridial Development (Text Figs. 26–39).
Fig.Figs. 26–33—Stages in development of the antheridium, × 735; Fig. 28—Anaphase, first meiotic division. Fig. 33—Successive sections of antheridium showing 32 nuclei at early anaphase in last nuclear division. Fig. 34—Tuft of antheridial paranemata, × 82. Fig. 35—Antheridia (fresh unstained), × 350; Fig. 36—Release of sperms from endochite, × 735. Fig. 37—Sperms (fresh), × 745. Fig. 38—Sperms (fixed and stained), × 625, Fig. 39—Development of stigma, × 745.
which are a few vacuoles. The stigma, a conspicuous orange bar on the yellow chromoplast, is terminal, borne on the anterior necklike portion of the sperm which usually projects from the rest of the body of the sperm, but sometimes appears to be bent back along the flattened side.
Stages in the development of the stigma as a modification of a plastid have been observed. It arises as a brilliant orange bar on a small plastid and enlarges as the plastid grows. In fresh material treated with chlor-zinc-iodine, the plastid appears green, and the stigma shows clearly as a dark green bar. Two long flagella are inserted laterally on the flattened side of the sperm. One is directed forward and the other backward. The anterior flagellum is shorter
than the posterior one and appears to act as the “rudder,” for it is usually directed straight forward and does not exhibit such lashing, whipping, coiling movements as does the more active posterior one. At times the sperm moves with such speed and contortions that the flagella are quite invisible. The length of the anterior flagellum averages 10 μ and the posterior flagellum, 13·5 μ These features have been determined from fresh material, confirmed from fixed and stained preparations.
The male gametes have a comparatively short life in suspension, but some may remain alive for several hours, their movements gradually becoming more sluggish, until finally they die. A large number give a distinctly orange tinge to a small quantity of Water, on account of the carotin present in the chromoplast and stigma of each sperm.
Fertilization occurs when active sperms are mixed with ova which have partially or completely lost their envelopes. This is best achieved in the laboratory by immersing a male receptacle or frond in a dish containing many ova. The sperms immediately
Fertilization—(Text Figs. 40–45).
Fig. 40—Sperm beside unfertilized ovum fixed 15 mins, after mixing male and female gametes, × 930. Figs. 41, 42 and 43—Passage ot male nucleus to female nucleus (from material fixed 30 mins, after mixing gametes), × 973. Fig. 44—Zygote one hour after mixing gametes. (Denser portion of reticulum may indicate male chromatin incompletely distributed in fusion nucleus), × 930. Fig. 45—First nuclear division in the zygote (from material fixed 38 hours after mixing gametes); spindle, centrosomes and asters conspicuous, × 973.
Fig. 5.—Photomicrograph of L.S. sporeling at 3 ½ mouths × 275, showing apical pit and apical paraphyses
Fig. 6.—Photomicrograph of L.S. young apical receptacle × 75, showing apical growing point. (Mucilage slightly displaced)
Fig 7.—Hormosira banksii forming a pure association of “hormosiretum,” Grove Arm, Queen Chailotte Sound, Manlborough
surround the ova in varying numbers. Each attacks the ovum, gyrating on the posterior flagellum with a “corkscrew” action, from time to time bunting against its surface. When many sperms attack one ovum the lashing of the flagella causes it to rotate spasmodically, sometimes quite fast, but very many sperms do not rotate the ovum, only moving it slightly to and fro. The rotation is merely an incidental, mechanical effect of the great activity of the flagella of the attacking sperms. Sperms also accumulate round immature ova and foreign bodies in the culture, so it cannot only be chemotaxis which accounts for their attraction to the mature unfertilized ova.
After a variable length of time, up to half an hour, one sperm penetrates into the ovum and the remaining sperms undergo a great change. Sometimes, when only a few sperms are present, the remainder swim away from the ovum, which no longer attracts and sperms. Usually, however, when many surround the ovum and one fertilizes it, the rest of the sperms become sluggish, as if affected by a toxic influence, and they soon die. Vast numbers of dead and dying sperms may be seen at the same time in a culture, but the mortality is not altogether simultaneous, as adjacent ova may be surrounded, one by active sperms, and the other by dead and dying sperms. After a short time all the sperms die in parts of the culture dish where ova have been fertilized. It is probable that the newly fertilized ova exert some influence over the sperms in their proximity, and this conclusion is strengthened by the fact that sperms released at the same time in water containing no ova remain alive long after those added to water containing ova.
If, five minutes after mixing ova and sperms, when only a few ova will have been fertilized, the sea-water medium is greatly diluted with fresh water, most of the ova swell and disintegrate diffusely, indicating that they were naked protoplasts. A few burst at one point indicating the presence of a bounding membrane. Ten minutes after mixing, similar treatment causes considerably more ova to burst at one point. It is evident that on fertilization, a membrane is immediately secreted by the zygote. At first it is very fine and only distinguishable by shrinking or swelling the enclosed protoplast, but later it thickens enough to be easily visible. (Text Fig. 49.)
After fertilization, other changes occur. The “granular,” finely alveolar, dense layer of cytoplasm round the central nucleus becomes more apparent and the phaeoplasts become orientated radially from the boundary of this layer. They form a dense aggregation occupying about one-half the diameter of the zygote, and extend radially between the large vacuoles of the “foamy” cytoplasm. In material fixed quarter of an hour after ova and sperms were mixed, sections of zygotes are easily distinguishable from those of unfertilized ova, even if the actual sperm is not visible, because of this characteristic approach to radial arrangement and aggregation of the phaeoplasts.
Stages in the passage of the male nucleus to the female nucleus within the ovum are shown in Text Figs. 41–43 from material fixed half an hour after ova and sperms were mixed. The central
large female nucleus usually has a single large nucleolus and a delicate, clearly-staining reticulum.
The sperm nucleus is oval or pear-shaped, and the individual chromosomes stain deeply, probably indicating that the nucleus of the sperm is at early telophase, persistent from the last nuclear division in the antheridium. By half an hour after mixing ova and sperms, the male nucleus has reached the periphery of the female nucleus. By one hour after mixing, most of the male nuclei have lost their identity in the fusion nuclei. Complete fusion is thus accomplished within one hour after the sperm has access to the ovum.
Germination. (Text Figs. 50A-J.)
Before germination takes place the fertilized ovum secretes a wide envelope of mucilage, which firmly attaches it to the substratum. Germination normally begins after a resting period of 18 to 24 hours in optimum conditions at room temperature, but it may be delayed until 40 to 48 hours after fertilization.
The polarity of the plant is first revealed by a protuberance from the zygote which may appear before or after the first division of the nucleus.
Conditions which may govern the orientation of the first cleavage and polarity in the Fucaceae have been described by various workers,
Germination (Text Figs. 46–51)
Fig. 46—Unfertilized ovum, × 225. Fig. 47—24 hours after fertilization, × 225, Figs. 48 and 49—Unfertilized ovum and zygote treated with chlor-zinc-Iodine, × 333. Fig 50—A.J. stages in germination present in culture three days after tertilization, × 225. (Semi-diagrammatic. Mucilaginous sheath not shown.) Fig 51—Typical sporeling at 4–6 days, × 250. (Mucilaginous sheath not shown.
and include: a directed beam of light (Farmer and Williams, 1898), especially the short blue light (Hurd, 1920); an electric current (Lund, 1923), the rhizoidal cell lying towards the positive pole; proximity, of neighbouring cells (Hurd, 1920); oxygen tension; products of metabolism, etc. For Hormosira, in the dark the first rhizoidal protuberance appears at random, but Hurd's “group effect” may be present. Zygotes situated within two or three diameters of one another tend to send out rhizoids towards each other or towards a nearby group.
Text Fig. 45 shows the first nuclear division in the germinating zygote, 38 hours after fertilization. Centrosomes, asters, and spindle are conspicuous, lying in an area of clear homogeneous cytoplasm around which the cytoplasm is denser and more vacuolate and contains many phaeoplasts.
Germination is not simultaneous in a culture, and material fixed at 38 hours contained stages from ungerminated zygotes to two-nucleate sporelings The daughter nuclei are surrounded by an aggregation of phaeoplasts, and the tip of the rhizoidal protuberance contains colourless cytoplasm at first. A cell-well may appear immediately after the first division, or it may not form until some time after the spindle has disappeared. It eventually divides the young sporeling transversely into dissimilar halves. The basal cell undergoes another transverse division which cuts off the rhizoid mother-cell. The other two cells give rise to the body of the sporeling, and the second transverse division may be preceded by a vertical division of the upper cell. Subsequent wall formation is variable, but the most typical form of young sporeling is shown in Text Fig. 51 (four and a-half days).
At all stages subsequent to germination the sporeling body and its rhizoids are surrounded by a continuous mucilaginous sheath.
During the first week, the first rhizoid elongates and becomes two or three-celled and up to 150 μ. long. Its tip is highly sensitive to light, and subjected to sudden unilateral illumination it grows away from the source of light, even turning through an angle of nearly 180°. Such negative phototropic response causes conspicuous orientation in one direction of all the rhizoids in a culture subjected to constant unilateral light. No such orientation takes place in horizontal cultures receiving light equally from all sides or only from above, or in cultures growing in the dark.
In some actively growing cultures the beginning of the second rhizoid was observed at the end of the first week. The cell at the proximal end of the first rhizoid divides longitudinally and/or transversely, and it is from this group of cells that later rhizoids arise. The second rhizoid appears as a protuberance, more usually after 10 to 12 days (Text Figs. 52, 53). Subsequent rhizoid-formation varies greatly, up to twelve having been counted, but three to six are most common.
By the end of the second week, the sporeling is 300–350 μ long. The two or three distal, elongated, rhizoidal cells have conspicuous spherical nuclei and some phaeoplasts in vacuolate cytoplasm. In the proximal rhizoidal region they are very numerous. Streaming
in the cytoplasm is easily seen in the large rhizoidal cells. The body of the sporeling has become darker in colour and in most cultures some of the sporelings grow upright in the dish.
By the end of the third week the sporeling body measures not less than 90 μ long, and has become more pear-shaped and less rounded at the apex.
Apical Paraphyses and Establishment of Apical Growth.
The first sign of the appearance of apical paraphyses is a slight flattening of the sporeling at the apex and the pressing apart of the cells between which the transparent tip of the first paraphysis protrudes (Text Figs. 54, 55). It pushes up the mucilaginous sheath and grows through it. Under optimum conditions the paraphyses may be quite long by the end of three weeks, but in some cultures they did not appear until after four weeks (especially in the winter months).
Later Sporeling Development (Text Figs. 52–62)
Figs. 62 and 53—Sporelings at 12 davit; Fig. 52—x 222, second rhizoid developing from proximal cell of first rhizoid. Fits. 53—x 120, second rhizoid elongated. Fig. 54 and 55—Sporelings at 4–5 weeks, × 148; first apical paraphysis showing. (P). Fig. 56. 57 and 58—Optical sections of sporelings at three months, × 55. Paraphysis in Fig. 58 measured 1.53 mm. (typical). Fig. 59—Sporeling at 6 months, × 55. No paraphyses remaining, sporeling body measured 173 μ lone, culture dish turned through 90°. Fig. 60—Optical section of dichotomous sporeling at 3 months, × 148. Figs. 61 and 62—Sections from paraffinager blocks; Fig. 61–L. S. sporeling at 7 ½ weeks, × 233. Fig. 62–L. S. sporeling at 3 ½ (Camera lucida drawing from two consecutive sections.)
They grow very long (as many as 30 cells), delicate, multicellular, unbranched hairs waving freely in the water (Text Figs. 56–58). After some months in culture they begin to break off and disintegrate until only a few cells project from the apex of the sporeling.
Longitudinal sections of the sporelings (Text Figs. 61, 62) show that the paraphyses grow from the floor of a depression or pit at the apex of the sporeling (somewhat similar to a very young cryptostoma). In Fucus vesiculosus (Neinburg, 1931), they are described as arising by periclinical division of some of the cells at the apex of the cylindrical sporeling (Delf, 1939).
At first the pit is not very large and the hairs fill the space as they grow out from the apex. The hairs grow by basipetal segmentation from, large, flask-shaped, basal cells, the products of which are small at first but which elongate distally. In older sporelings, the cavity is more spacious and contains some mucilage. Delf (1939) records that Neinburg (1931) found in Fucus vesiculosus the basal cell of one of the most central of an apical group of hairs finally gave rise to the apical cell of the future thallus. He believed that the single apical cell was formed by inclined walls within the special hair cell, but this was observed only in hairs formed in connection with regeneration following injury to the thallus.
I have cut sections of sporelings up to 4 ½ months old, but there has not been conclusive evidence of what occurs in sporelings of Hormosira banksii to give rise to the four apical cells of the future thallus. I have not found more than three hairs projecting from the apex of any sporeling, but in some sections an undeveloped fourth hair was present as a short column of cells. Whether this is of common occurrence, and the four basal cells eventually give rise to the four apical growing-cells, has not been ascertained. It is possible that one of the large basal cells of the hairs undergoes two divisions to form the four apical cells, and the other cells disintegrate, as in Fucus.
Differentiation of the tissues of the young plant takes place after the establishment of apical growth.
Dichotomy.—Some sporelings exhibit an early dichotomy (Text Fig. 60). They grow considerably wider anteriorly instead of the usual pear-shape, and two apical pits form, from each of which one, two, or three paraphyses grow in the same way as for the single apex.
Culture of Six Months.—In laboratory cultures little apparent growth occurs after about four months, though the sporelings in a culture of six months appear to be still alive. They are dark brown in colour, cylindrical or pear-shaped, and are easily visible to the naked eye. The rhizoids are very long (up to 1 8 mm.) and the sporeling body measures up to 180 μ in length (Text Fig. 59).
Alternation of Generations.
The vegetative thallus of Hormosira is diploid, the haploid phase being confined so the sex organs. Meiosis occurs in the first two divisions in the sex organs, followed by one division in the
oogonium and by four divisions in the antheridium. The diploid condition is re-established on fertilization. Gametic-zygotic numbers of 12–24, have been tentatively determined from chromosome counts of antheridial stages, sperm nuclei, and vegetative nuclear divisions.
The brown alga Hormosira banskii (Turner) Decaisne is a monotypic, Australasian genus belonging to the group Anomalae of the family Fucaceae.* It inhabits rock platforms and pools between tidemarks. Internal water reserves in the hollow receptacles make it, to a large extent, drought resisting.
Growth is from an apical group of four large tetrahedral cells.
The cryptostomata are all potential sexual conceptacles, and the basal cell derived from the initial cell gives rise to the greater part of the floor and walls, supplemented by cells from the “limiting” layer or meristoderm.
Hormosira is dioecious and fertile all the year round. The oogonium contains four functional ova and undeveloped ova may persist and be released with the female gametes.
Stages in the development of the antheridum are described. The stigma of the sperm is formed by modification of a plastid.
Fertilization may be accompanied by rotation of the ovum by the sperms, which die in proximity to newly fertilized ova. The passage of the male nucleus to the female nucleus within the ovum is described from sectioned material. The fusion nucleus is formed within one hour after the sperm has access to the ovum.
Preliminary stages of the origin of the apical growing point in the young sporeling are described.
The vegetative thallus is diploid and the haploid stage is confined to the sex organs, the diploid complement being restored on fertilization. The gametic-zygotic numbers are tentatively determined as 12–24.
I wish to acknowledge my indebtedness, for valuable encouragement and suggestions, to Dr. I. V. Newman, Victoria University College, Wellington, New Zealand.
Agardh, J. G., 1848. Species Algarum, vol. 1, p. 108.
—— 1877. De Algis Novae Zelandiae Marinis. Lunds Univers. Årsskrift, T. 14.
Barton, E. S., 1899. On Notheia anomala Harv. et Bail. Jour. Linn. Soc. Botany, 34, p. 417.
Belling, 1921. In McClung, Microscopical Technique, p. 165, 1937.
Bory, 1827. Voyage autour le monde—La Coquille 1822–25.
Bower, F. O., 1880. On the Development of the Conceptacle of the Fucaceae. Quart. Jour. Micro. Science, p. 30.
Chamberlain, C. J., 1932. Methods in Plant Histology, p. 27.
Decaisne, 1842. Classif. des Algues. Ann. de Sci. Nat. Bot., ser. II, 17, p. 330.
Delf, M., 1939. The Systematic Position of the Fucales. The New Phytologist, 37, No. 3, p. 224.
de Toni, J. B., 1895. Sylloge Algarum, vol. 3, p. 188.
Endlicher, 1836. Genera Plantarum, p. 10; supplement 3, p. 29. (Reference supplied by Mr. R. M. Laing, Volume not available in New Zealand.)
Farmer, J., and Williams, J. L. 1896–7. On Fertilization and Segmentation of the Spore in Fucus. Proc. Roy. Soc. Lond., 60, p. 193.
[Footnote] * See footnote p. 2.
Farmer, J., and Williams, J. L., 1898. Contribution to our knowledge of the Fucaceae. Roy. Soc. Lond. Trans. B, 190, p. 623.
Fritsch, F. E., 1935. The Structure and Reproduction, of the Algae, vol. 1, p. 23.
—— 1945. The Structure and Reproduction of the Algae, vol. 2.
Getman, M. R., 1914. Oogenesis in Hormosira. Bot. Gaz., 58, p. 204.
Gibb, 1937. Himanthalia lorea. Jour. Linn. Soc. Botany, 51, no. 337.
Gruber, 1896–97. Uber Aufbau und Entwicklung einiger Fucaceen. Bibliotheca Bot., 38.
Harvey, W. H., 1860. Phycologia Australica, vol. 3.
—— 1867. In Hooker's Handbook of the New Zealand Flora′
Harvey and Bailey, 1802. United States Exploring Expedition (Capt. Wilkes), 18, Botany, p. 157. Reference quoted in Barton above.
Hedley, C., 1915. An Ecological Sketch of the Sydney Beaches. Pres. Addr., Roy. Soc. N.S.W., 49, p. 57.
Hick, T., 1885. Protoplasmic Continuity in the Fucaceae. J. Bot. N.S., 14, p. 97, (Delf, 1939).
Hooker, J. D., 1867. Handbook of the New Zealand Flora. Seaweeds by W. H. Harvey.
Hurd, 1920. From Whitaker, 1931.
Kutzing, F. T., 1849. Species Algarum.
Laing, R. M., 1885. Observations on the Fucoidea of Banks Peninsula. Trans. N.Z. Inst., 17, p. 303.
—— 1899. Revised List of New Zealand Seaweeds, Part 1. Trans. N.Z. Inst., 32, p. 37.
—— 1921. Island Bay, Wellington, a Collecting-ground for Marine Algal Vegetation. N.Z. Jour. of Sci. and Technology., vol. 4, no. 4, p. 204.
—— 1926. A Reference List of New Zealand Marine Algae. Trans. N.Z. Inst., 57, p. 120.
—— 1928. The External Distribution of the New Zealand Marine Algae and Notes on some Algological Problems. Trans. N.Z. Inst., 58, p. 189.
Lesson, A., et Richard, A., 1832. Voyage de L'Astrolabe 1826–29 sous le commandement de M. Dumont D'Urville; Botanique per Lesson et Richard. (Seaweeds by Richard.)
Lucas, A. H. S., 1909. Revised List of the Fucoideae and Florideae of Australia. Proc. Linn. Soc. N.S.W., 34, p. 1.
—— 1913. Notes on Australian Marine Algae. Trans. Linn. Soc. N.S.W., vol. 38, p. 51
Lund, 1923. From Whitaker, 1931.
Madge, 1936 The Use of Agai in Embedding. Ann. Bot., vol. 50, 199, p. 677
May, V., 1939. Ectocarpus confervoides (Roth.) le Jol Proc. Linn. Soc. N.S.W., vol. 64, p. 544.
—— 1939. Key to the Marine Algae of N.S.W. Trans. Linn. Soc. N.S.W., vol. 64.
McClung, C. E., 1937. Microscopical Technique, p. 165.
Mitchell, M. O., 1893. Notheia anomala. Phycological Memoires.
Mollett, T. A., 1880. On the Structure of Hormosira billardieri. Trans. N.S. Inst., 13, p. 318.
Montagne, 1845. Voyage au Pole Sud, Bot, Crypt.
Nienburg, W., 1931. Die Entwicklung der Keimung von Fucus vesiculosus und ihre Bedeutung fui die Phylogenie der Phaeophyceen. Wiss. Meeresunter. Abt. Kiel, 21, p. 51. (Reference quoted from Delf, 1939. Volume unobtainable in New Zealand or Australia.)
Oliver, W. R. B., 1923. Marine Littoral Plant and Animal Communities in New Zealand. Trans. N.Z. Inst., 54, p. 490.
Oltmanns, F., 1922–23. Morphologie und Biologie der Algen, vol. 2.
Parke, M., 1933. A Contribution to the Knowledge of the Mesogloiaceae and Associated Families. Publ. Hartley Bot. Lab. L'pool Univ., no. 9 (Delf, 1939).
Rees, Ethel M., 1933. Observations on Bifurcaria tuberculata Stackh. Ann. Bot. Lond., 185, 101 (Delf, 1939).
Roe, M. L., 1916. The Development of the Conceptacle in fucus. Bol. Gaz., vol. 61, p. 231.
Turner, 1808. Fuci sivi Plantanum Fucorum Generi a Botanicus Ascriptarum. Historica Fucorum, vol. i. p. 1.
Whitaker, 1931. Biological Bulletin, p. 294.
Williams, M., 1923. Contribution to our Knowledge of the Fucaceae. Trans. and Proc. Linn. Soc. N.S.W., vol. 48, p. 634.