Microplankton analysis

2.4.1. Sample preparation. Two aliquots from Lugol’s fixed initial samples and three random samples from experimental and control bottles were analysed from each 24 h incubation period. Each sample bottle was carefully rotated through 360 degrees at least 50 times to ensure that all matter was re-suspended and fully mixed. The samples were initially concentrated in parafilm-sealed, 100 ml measuring cylinders. Entire samples (100 ml) were settled from the spring cruise (D262), whilst only half the sample volume (50 ml) of the Summer Cruise (D264) samples was settled. After the time required to ensure complete sedimentation (24 and 48 h for 50 and 100 ml respectively: Lund et al. 1958, Gifford 1993, Gifford and Caron 2000), the supernatant water was slowly and carefully removed to a clean storage bottle until approximately 20 ml of sample remained. Cells were then re-suspended by rotating the cylinder between the palms of the hands for 30 seconds (see Lund et al. 1958) and transferred to a 25 ml settling chamber (Duncan and Associates: http://www.duncanandassociates.co.uk/).

Because all phytoplankton samples were preserved with 10 % acid Lugol’s solution, they required a degree of bleaching (removal of iodine) before accurate identification of the cells could be made. This was achieved by carefully adding drops of a saturated sodium thiosulphate solution (in milli-Q water) to the sample in the 25 ml settling chamber (Sherr and Sherr 1993). The bleaching technique, initially tested on samples from the ‘trial cruise’ D260, typically required 4 drops of sodium thiosulphate to turn the sample clear. The remaining chamber volume (~4ml) was topped up with the supernatant water (10 % Lugols vol:vol), returning the sample to a ‘weak tea’ coloured solution, before applying the glass coverslip. Over-addition of the bleaching solution caused the thiosulphate to crystallise on the baseplate of the chamber, and in severe cases, completely obscured the sample. In cases of over- addition, neat Lugol’s was dripped into the chamber until the familiar ‘weak tea’ colour was achieved. Following a final 12 h period of sedimentation, the cells were then enumerated by means of inverted microscopy, the protocol of which is described by Lund et al. (1958).

2.4.2. Cell counts. Cell counts were undertaken on an Olympus IMT-2 inverted microscope in a darkened room. All cells excluding flagellates and

cryptomonads were enumerated at X 200. Flagellates and cryptomonads (all < 10µm)

were counted at X 400 on a single ‘field of view’ transect from top to bottom using phase contrast. The area of the flagellate transect was determined as diameter of baseplate (23 mm) multiplied by the width of the field of view (0.048 mm). By

expressing this area (1.104 mm2) as a fraction of the entire baseplate area (415.48

mm2), the multiplication factor of 376.341 is calculated (415.48/1.104). By assuming

that the distribution of flagellates within this single transect was representative of the distribution of flagellates throughout the baseplate, the number of flagellates per volume of sample settled was calculated by applying the multiplication factor to the number of flagellates counted in one transect. In the majority of cases > > 100 cells

were counted, providing a 95 % confidence interval of the estimate within ± 20 % x

(Lund et al. 1958, Venrick 1978).

2.4.3. Reliability of the cell counts. Typically >> 100 cells for each individual group were counted. As discussed by Venrick (1978 and refs therein), counting 100 cells is sufficient to give a 95 % confidence interval of the estimate

within ± 20 % x. Before undertaking any sample analysis, each cell group in 8

samples from experiment 1, D262, were counted, then re-counted and the results statistically compared. So as not to influence the latter counts by the previous ones, group counts were only summed after both counts had been completed. Randomization was achieved by the physical mixing of the samples before settlement (Venrick 1978). It is therefore valid to compare two single sample counts (Parker 1983): ] 4 / ) [( 5 . 0 ] 2 / ) ( [ 2 1 2 1 1 X X X X X d + − + − = (1)

where X1 and X2 are the two counts and d is the ‘standardized normal deviate’ (∞).

In all cases, the counts were not significantly different from each other (p > 0.1 in all cases).

2.4.4. Cell volume estimations. For each defined group, the appropriate linear measurements of at least 30 fixed cells were made with a calibrated graticule

in the ocular of the Olympus IMT-2 inverted microscope. Cell volumes were estimated using simple geometric formulae (Table 2.1), as suggested by Menden- Deuer and Lessard (2000).

2.4.5. Volume:Carbon regression equations. Strathmann (1967) made the important distinction between the cell volume to carbon (vol:C) relationships for diatoms and for other protists (because of their large vacuoles, diatoms are less carbon dense), demonstrating the need for separate predictive equations. Despite being adopted as a standard method (e.g. Parsons et al. 1984), little attempt has been made to justify the use of these equations. Considering that the cellular C of cultured organisms (typically used in determining conversion factors) is influenced by the culture conditions (Putt and Stoecker 1989, Thompson et al. 1991, 1992, Davidson et al. 2002), and that the relatively few cultured organisms used are rarely the same as those encountered in field based studies, this is somewhat surprising.

Vol:C conversion factors have subsequently been determined for various components of the microplankton, including phototrophic nanoplankton (Verity et al. 1992), flagellates (Borsheim and Bratbak 1987), dinoflagellates (Menden-Deuer and Lessard 2000), ciliates (Putt and Stoecker 1989), diatoms (Strathmann 1967) and various phytoplankton (Mullin et al. 1966; Montagnes et al. 1994). More recently, Menden-Deuer and Lessard (2000) determined highly significant vol:C relationships for marine protists using both new experimental work and all existing data in the literature. The C biomass of diatoms and protists excluding ciliates (see below) was estimated here using the corresponding equations presented by Menden-Deuer and Lessard (2000).

The C density of aloricate (naked) ciliates is on average 43 % more dense than similar sized dinoflagellates (Menden-Deuer and Lessard 2000). Accordingly, aloricate ciliate C biomass is calculated using Putt and Stoecker’s (1989) regression for 2 % acid Lugols preserved aloricate ciliates.

2.4.6. Shrinkage effects of Lugol’s. Before C biomass was calculated, the cell volumes of all non-ciliate and non-thecate dinoflagellate taxa were adjusted to account for shrinkage due to preservation with acid Lugol’s (Appendix 1). Due to the uncertainties in predicting preservation-induced cell volume changes in thecate dinoflagellates (Menden-Deuer et al. 2001), their volume was not corrected.

Table 2.1. Geometric formulae used to estimate cell volume, where L is length (the longest straight line separation between any two points on the cell boundary regardless of orientation), B is breadth (widest distance measured perpendicular to

length), H is height and R is radius (i.e. B/2). §H is determined by an aspect ratio of

0.5 * B

Cell group Shape

approximation Formula

Dinoflagelates

(naked and thecate) Prolate spheroid (Pi/6)*L*B^2

Nitzschia spp. Two pyramids (1/3*(B^2)*(L/2))*2

Pennate diatom

(Triponeis sp.) Cylinder Pi*R^2*L

Centric diatom Cylinder Pi*R^2*H§

Ciliate Prolate spheroid (Pi/6)*L*B^2

Silicoflagelate Sphere (Pi/6)*L^3

Flagellate < 3.5 µm Sphere (Pi/6)*L^3

Flagellate > 3.5 µm Sphere (Pi/6)*L^3