Journal of Applied Phycology 11: 285–291, 1999. © 1999 Kluwer Academic Publishers. Printed in the Netherlands.

285

The relative sensitivity of algae to decomposing barley straw Derek Martin∗ & Irene Ridge Department of Biology, Open University, Walton Hall, Milton Keynes MK7 6AA, UK (∗ Author for correspondence) Received 14 March 1999; revised and accepted 31 March 1999

Key words: barley straw, inhibition, stimulation, diatoms, Cyanophyta, Euglena gracilis

Abstract Decomposing barley straw has previously been shown to inhibit the growth of a limited number of algae under both laboratory and field conditions. Bioassays were conducted on a range of algae to evaluate their relative sensitivities to straw-derived inhibitor(s). A range of sensitivities was found, including some species that were resistant to the straw-derived inhibitor(s). A microcystin-producing strain of Microcystis aeruginosa was very susceptible to decomposing barley straw. Bioassays using Euglena gracilis suggest that the inhibitory compounds are not derived from the phototransformation of straw decomposition products and do not act primarily by inhibiting photosynthesis. Susceptibility to barley straw appears not to be related to general taxonomic or structural features. Possible implications for algal populations in natural freshwaters are briefly discussed.

Introduction Welch et al. (1990) reported the algal-inhibiting properties of barley straw (Hordeum vulgare) when decomposed in water in the field. Subsequent research suggested that the inhibitory component derived from the straw itself and not from the associated mycoflora, although the production of algal antibiotics by bacteria was not ruled out (Pillinger et al., 1992). Pillinger et al. (1994) proposed the main algal-inhibiting factor(s) to be some form of oxidized polyphenolic compound derived from lignin which was solubilized from the straw. Laboratory studies suggest that the inhibitor(s) are algistatic, rather than algicidal (Gibson et al., 1990; Newman & Barrett, 1993). Barley straw is now in widespread use as a method of algal control since it is relatively inexpensive and no adverse ecological effects have been reported. The growth of macrophytes and animal life appear unaffected (Everall & Lees, 1996) and barley straw has been used in potable water supplies, where no unusual amounts of the organic chemicals routinely monitored by the water industry have been found (Barrett et al., 1996).

A number of other materials have also been found to be anti-algal including brown-rotted wood (Pillinger et al., 1995; Ridge & Pillinger, 1996) and some leaf litters, in particular oak leaves (Quercus robur) (Ridge et al., 1995). In the case of oak leaf litter, oxidized tannins leached during the first stages of decomposition may be the initial source of algal inhibition, but the continued inhibition of algal growth over many months suggests that lignin degradation is the source of the inhibitor(s) in leaves and barley straw alike. It has been suggested that diatoms appear to be resistant to the inhibitor(s), since one species was found to grow in tanks of decomposing barley straw (Ridge et al., 1995). However, in field trials in a reservoir, Barrett et al. (1996) found that the planktonic diatoms Asterionella formosa and Tabellaria sp. were suppressed after the introduction of barley straw, although no truly comparable control was used in this study and cell counts had to be compared to those obtained from the reservoir in previous years. Desmids have also been observed in ponds which had been treated with barley straw or oak leaves, where the growth of Clado-

286 phora glomerata was shown to be inhibited (authors’ unpublished data). In the laboratory Microcystis aeruginosa was shown to be inhibited with as little as 2.57 g dry mass of barley straw m−3 growth medium (Newman & Barrett, 1993) and Gibson et al. (1990) showed that a number of green algae were inhibited, but with doses of up to 104 g dry mass m−3 growth medium. In the field, effective algal control can be achieved with as little as 3–5 g dry mass m−3 water although doses of 20–50 g dry mass m−3 water provide a wider safety margin (Ridge et al., 1995). The evidence suggests that there are differences in susceptibility between taxa of algae, hence, this study was undertaken to identify the relative sensitivity of a range of taxa to decomposing barley straw. Comparisons of the differing sensitivity/resistance of a range of taxa could give clues as to the mode of action of the inhibitor(s) from the underlying physiology and morphology of the species in question. Test species were chosen because they were known to be able to grow to nuisance proportions, or for their ability to tolerate stresses such as high temperatures and desiccation, or where there were interesting structural features which may have affected sensitivity towards straw-derived inhibitor(s) (e.g. siliceous cell wall of diatoms). Materials and methods Culture media and algal strains The growth media used are shown in Table 1. A single growth medium was not adopted, since media were chosen to obtain the best possible growth of control cultures. The test species were grown in mineral media, apart from Euglena gracilis which was grown in an organic medium that supported heterotrophic nutrition. All stock cultures were maintained by continuous sub-culture and both the stock and experimental growth media were buffered with 20 mM 4-(2-hydroxyethyl) piperazine-1-ethane sulphonic acid sodium salt. All of the stock cultures were unialgal but no attempt was made to maintain them axenically. The JM1 medium was based upon the JM medium of Tompkins et al. (1995) modified by Newman & Barrett (1993) and buffered to pH 8.2. The WC medium was after Guillard & Lorenzen (1972) and buffered to pH 7.2. Both the DM and EG media were after Tompkins et al. (1995) and buffered to pH 8.0. The origin of the strains is shown in Table 1. CCAP strains were obtained from the Culture Col-

lection of Algae and Protozoa, Ambleside, UK; SAG strains were obtained from Sammlung von Algenkulturen, Göttingen, Germany and Sciento strains were obtained from Sciento, Manchester, UK. Stichococcus bacillaris was obtained from the culture collection of the Natural History Museum, UK. Tabellaria flocculosa was isolated from Loch nam Brac, Scotland (58◦ 230 N, 5◦ 60 W) and the Loe Pool strain of Microcystis aeruginosa was isolated from Loe Pool, a eutrophic lake in Cornwall, UK (50◦ 40 N, 5◦ 170 W). The Loe Pool strain was assayed after only one sub-culture in the laboratory, while it still retained its colonial form. The other strains of M. aeruginosa tested no longer retained their colonial form and grew as single cells. The AK1 strain of Microcystis aeruginosa was supplied by Prof. G. A. Codd (University of Dundee) and it produced microcystin in culture at approximately 0.2 µg microcystin-LR equivalents mg−1 dry mass (G. A. Codd, pers. comm.). Nitzschia filiformis var. conferta was isolated from a tank of oak leaves which had been decomposing in the light. The Synechococcus sp. was isolated from contaminated growth medium, which had been accidentally heated to approximately 35 ◦ C. Preparation of straw and bioassay Active barley straw was obtained by incubating 2000 g dry mass m−3 in aged tap water, in darkness, at room temperature for three to six months with vigorous aeration (Pillinger et al., 1994). All bioassays were performed in 150-mL conical flasks. Wet straw was cut into pieces ≤ 5 mm in length, placed into 50 mL of the relevant medium and then inoculated with 1 mL of an exponentially growing culture of the test species. The dry mass of straw was found by drying over silica gel, under reduced pressure, until mass was constant. The mean mass of dry straw was ca 10% of wet mass. All flasks were placed in a growth cabinet (SANYO-Gallenkamp) at 20 ◦ C ± 1 ◦ C lit from above by fluorescent tubes with continuous light of 120 µmol m−2 s−1 PAR and shaken twice daily. Euglena gracilis was grown both in the dark (heterotrophic growth) and the light (photoheterotrophic growth). The duration of the assays is shown in Table 1. Biomass estimation Wherever possible the yield of algae at the end of an assay was quantified by cell counting on a haemacytometer or Sedgewick-Rafter chamber (Table 1). Where

287 Table 1. Origin and bioassay conditions of the test species. Phylum

Species

Origin

Growth medium

Biomass estimation method

Duration of assay (days)

Bacillariophyta

Asterionella formosa Tabellaria flocculosa Nitzschia filiformis var. conferta

Sciento Loch nam Brac decomposing oak leaves

DM DM X3 DM

count count count

4 5 4

Chlorophyta

Closterium ehrenbergii Staurastrum pingue Cosmarium biretrum Spirotaenia erythrocephala Chlorella vulgaris Scenedesmus subspicatus Pediastrum boryanum Stichococcus bacillaris

SAG 134.80 SAG 5.94 SAG 44.86 SAG 7.89 CCAP 211/12 CCAP 276/20 Sciento BM 83092907A

WC WC WC WC JM1 JM1 JM1 JM1

count count count chl a count count count count

8 7 7 8 4 4 4 5

Euglenophyta

Euglena gracilis

Sciento

EG

count

6

Cyanophyta

Anabaena flos-aquae Anabaena cylindrica Aphanizomenon flos-aquae Oscillatoria redekei Oscillatoria animalis Microcystis aeruginosa

CCAP 1403/13B Sciento CCAP 1401/1 CCAP 1459/29 Sciento Loe Pool CCAP 1450/6 Sciento AK1

JM1 JM1 JM1 JM1 JM1 JM1 JM1 JM1 JM1

chl a chl a chl a chl a chl a chl a count count count

8 8 8 8 8 4 4 4 4

Synechococcus sp.

contaminant

JM1

chl a

4

cell counting was made difficult by the size of cells, or where the algae were filamentous, chlorophyll a analysis was carried out according to the method of Pillinger et al. (1994). Briefly, at the end of an experiment the contents of each flask where filtered through a glass fibre filter (GF/C Whatman) and the filter plus residue were extracted with 100% methanol. Chlorophyll a was quantified by measuring the absorbance at 665 nm minus a background turbidity reading at 750 nm on a SP6-550 UV/VIS spectrophotometer (Pye Unicam) on both raw extract and acidified extract, to correct for phaeopigments (Marker et al., 1980). Experiments were performed on a range of doses of barley straw, using five replicate flasks for each dose, and an analysis of variance was performed to test for significance. The results are expressed as the

mean dry mass of straw, of at least two replicate experiments, required to obtain a 50% reduction in yield when compared to a control value. Although the mass of straw which was employed in the bioassays could be controlled, the exact amount of inhibitor(s) and/or its rate of release could not be controlled, since this depended on the microflora which developed on the straw and the length of the decomposition period. For this reason the test species were categorized according to their sensitivities (Table 2). This allowed comparisons to be made without relying on exact 50% inhibition of yield values since they were subject to slight variations caused by variability in the straw.

288 Table 2. Susceptibility of algae to decomposing barley straw. Category

Species/strain

Barley straw required for 50% inhibition of yield (dry mass g m−3 medium)

Very susceptible 0–1000

Microcystis aeruginosa Sciento Loe pool AK1 CCAP 2301450/6 Tabellaria flocculosa Anabaena flos-aquae Closterium ehrenbergii Cosmarium biretrum Asterionella formosa Oscillatoria redekei Spirotaenia erythrocephala Synechococcus sp Staurastrum pingue

70 90 180 230 295 375 450 540 667 670 750 750 960

Susceptible 1001–2000

Aphanizomenon flos-aquae Pediastrum boryanum Stichococcus bacillaris

1370 1500 1940

Slightly susceptible > 2001

Euglena gracilis Chlorella vulgaris

Resistant (stimulation)

Nitzschia filiformis var. conferta Scenedesmus subspicatus Anabaena cylindrica Oscillatoria animalis

> 27001 > 5400 – – – –

1 Result obtained from Euglena gracilis grown in the light

Results The amount of straw required to obtain a 50% reduction in yield for all of the species which were assayed is shown in Table 2; in all cases inhibition was significant (p < 0.001). Barley straw inhibited the two planktonic diatoms Asterionella formosa and Tabellaria flocculosa, whereas the surface-associated diatom Nitzschia filiformis var. conferta was not inhibited and growth was actually stimulated. Placoderm desmids (Cosmarium biretrum, Closterium ehrenbergii and Staurastrum pingue) and a saccoderm desmid (Spirotaenia erythrocephala) were very susceptible to barley straw. Members of the Chlorococcales were found in all of the categories presented in Table 2 apart

from the very susceptible category. Cyanophyta were found to be both susceptible and resistant, although four strains of Microcystis aeruginosa were found to be the most susceptible of all the species tested. Growth of all the resistant species was stimulated in the presence of decomposing barley straw. In all cases stimulation was at least 200% of the control with up to 4000 g dry mass barley straw m−3 medium (p < 0.001). Euglena gracilis was only slightly susceptible to barley straw, but susceptibility varied depending on whether the cultures were grown in the light or the dark (Table 3). After three days growth the lightgrown cultures were not inhibited whereas those that were grown in the dark were inhibited in the pres-

289 Table 3. Yield of Euglena gracilis grown with decomposing barley straw (2727 g dry mass m−3 medium) in darkness and in light. Values are means ± S.E. (n = 5). Values for light and dark assays are compared to controls grown in the light and dark respectively (significant differences from the control (100%) are shown as ∗ p < 0.01, ∗∗ p < 0.001). Time (days)

Yield (% control) Light Dark

3 6

96 ± 8 67 ± 3∗

56 ± 6∗ 29 ± 8∗∗

ence of barley straw. After six days growth both the light- and dark-grown cultures were inhibited, although the dark-grown cultures were inhibited more strongly. The same pattern of inhibition was obtained in three replicate experiments.

Discussion Decomposing barley straw inhibited the growth of planktonic diatoms under the specified laboratory conditions. This is contrary to the report of Ridge et al. (1995) which suggested that diatoms as a group may not be inhibited. The present study supports the field observations of Barrett et al. (1996), who showed that Asterionella formosa and Tabellaria sp. were inhibited when barley straw was introduced into a reservoir. N. filiformis was not inhibited, as was perhaps to be expected, since it was isolated originally from a tank of decomposing oak leaves. Oak leaves have inhibitory effects comparable to those of decomposing barley straw (Ridge et al., 1995). Newman and Barrett (1993) suggested that Cyanophyta were particularly susceptible to decomposing barley straw, but the results presented in Table 2 do not support this view. Although all of the Microcystis aeruginosa strains were the most susceptible tested, both Anabaena cylindrica and Oscillatoria animalis were resistant. The AK1 strain of M. aeruginosa was very susceptible which, apparently, is the first time a known microcystin-producing strain has been shown to be susceptible to barley straw in the laboratory or in the field. The Loe Pool strain of M. aeruginosa, which was tested while it still retained its colonial growth form, was also very susceptible. Thus, the mucilaginous

mass which makes up a colony of M. aeruginosa gives no greater protection from straw-derived inhibitor(s) than that of single-celled forms of M. aeruginosa. The fact that inhibition of Euglena gracilis occurs at all in the dark shows that the phototransformation of straw decomposition products into phytotoxic compounds, proposed by Barrett (1994), cannot account for the inhibition observed in laboratory assays. The current results with E. gracilis support those of Cooper et al. (1997) who showed several species of Saprolegnia to be inhibited by barley straw when grown in the dark. Saprolegnia was formerly classified in the fungal class Oomycetes (Webster, 1980), but their characteristic heterokont zoospores suggest that they are more closely related to ’algae’ (van den Hoek et al., 1995). Although the straw-derived inhibitor(s) appear not to be acting on a photosynthetic process in E. gracilis, further work is required to determine whether photosynthesis can be discounted as the primary site of action for the straw-derived inhibitor(s) in other phototrophs. What is interesting is that E. gracilis shows a delayed and weaker inhibition when it is grown in the light rather than in the dark (Table 3). The ability to grow photoheterotrophically may increase resistance to some toxins, as reported by Megharaj et al. (1992), who showed that two species of Chlorococcales were resistant to certain phenolic compounds when growing photoheterotrophically, but not when growing auto- or heterotrophically. Further experiments using axenic cultures may be valuable to determine more clearly the growth characteristics of E. gracilis and to see if light confers some degree of resistance over longer periods of time. However, in the current assays the presence of decomposer organisms was a prerequisite for inhibitor release so, unless the straw is chemically treated to allow abiotic inhibitor release (Pillinger, 1993), the use of axenic cultures is problematical. Although only one of the desmids tested was known to form dense populations (Staurastrum pingue), other desmids were tested to determine whether differences in their cell wall structure (Brook, 1981; Gerrath, 1993) could affect sensitivity to barley straw. However, the results presented in Table 2 show that all of the desmids were very susceptible to barley straw. The concentrations of barley straw required for algal inhibition in the present laboratory study are larger than those which are reported to be necessary in the field (Ridge et al., 1995; Barrett et al., 1996). Geyer et al. (1985) suggested from work by Jou-

290 any et al. (1983) that organic chemicals would be more toxic under the dynamic conditions that occur in the field; a static test would underestimate toxicity. Whether this is true with the proposed chemicals released from barley straw requires further investigation. The difference in dosage highlights the problems of extrapolating these laboratory investigations to the field. However, of the small number of species which have been found to be susceptible in field studies (in reservoirs), Asterionella formosa, Tabellaria sp. (Barrett et al., 1996) and Aphanizomenon flos-aquae (Everall & Lees, 1996) were all susceptible in the current laboratory studies (Table 2). All of the resistant of species were stimulated by decomposing barley straw. Growth stimulation could be due to the presence of some unknown organic nutrient or perhaps certain organic substances originating from the barley straw are acting in a growth regulatory capacity (Larson, 1978). The inhibitory action is thought to be based on the same principle in barley straw, leaf litter (Ridge et al., 1995) and brown-rotted wood (Pillinger et al., 1995). The use of decaying plant litter (particularly barley straw) to control nuisance algae may have important implications from a water management point of view, but of possibly greater significance is the importance of these materials in natural ecosystems. Leaf litter and decaying wood are major inputs into freshwater ecosystems (Webster & Benfield, 1986) and could influence the species composition and population growth of algal communities if conditions were favourable. The laboratory studies presented here and by Gibson et al. (1990) suggest that the sensitivity of algae to decomposing barley straw is not related to general taxonomic or structural characteristics; even members of the same genus can differ widely in their sensitivity. The fact that some species appear to be resistant to barley straw may have important implications for water management, although no known nuisance species were shown to be resistant in this study. Further evidence is required to see whether these findings can be extrapolated into the field where natural inputs of plant litter may play a part in controlling algal biomass and/or species composition.

Acknowledgements We thank Dr E. J. Cox for identification of Nitzschia filiformis var. conferta, Dr B. A. Whitton for helpful

advice during manuscript preparation and the Open University Research Committee for financial support.

References Barrett PRF (1994) Field and laboratory experiments on the effects of barley straw on algae. In Hewitt HG (ed.), BCPC monograph 59: Comparing field and glasshouse pesticide performance II. British Crop Protection Council, Farnham: 191–200. Barrett PRF, Curnow J, Littlejohn JW (1996) The control of diatom and cyanobacterial blooms in reservoirs using barley straw. Hydrobiologia 340: 307–311. Brook AJ (1981) The Biology of Desmids. Botanical Monographs 16. Blackwell, Oxford, 276 pp. Cooper JA, Pillinger JM, Ridge I (1997) Barley straw inhibits growth of some aquatic saprolegniaceous fungi. Aquaculture 156: 157–163. Everall NC, Lees DR (1996) The use of barley straw to control general and blue-green algal growth in a Derbyshire reservoir. Wat. Res. 30: 269–276. Gerrath JF (1993) The biology of desmids: a decade of progress. In Round FE, Chapman DJ (eds), Progress in Phycological Research 9. Biopress, Bristol: 79–192. Geyer H, Scheunent I, Karte F (1985) The effects of organic environmental chemicals on the growth of the alga Scenedesmus subspicatus: a contribution to environmental biology. Chemosphere 14: 1355–1369. Gibson MT, Welch IM, Barrett PRF, Ridge I (1990) Barley straw as an inhibitor of algal growth II: laboratory studies. J. appl. Phycol. 2: 241–248. Guillard RRL, Lorenzen CJ (1972) Yellow-green algae with chlorophyllide C1,2 . J. Phycol. 8: 10–14. Jouany JM, Ferard J, Vasseur P, Gea J, Truhaut R, Rast C (1983) Interest of dynamic tests in acute ecotoxicity assessment in algae. Ecotox. envir. Saf. 7: 216–228. Larson RA (1978) Dissolved organic matter of a low coloured stream. Freshwat. Biol. 8: 91–104. Marker AFH, Nusch EA, Rai H, Reimann B (1980) The measurement of photosynthetic pigments in freshwaters and standardization of methods: Conclusions and recommendations. Arch. Hydrobiol. Beih. Ergebn. Limnol. 14: 91–106. Megharaj M, Pearson HW, Venkateswarlu K (1992) Effects of phenolic compounds on growth and metabolic activities of Chlorella vulgaris and Scenedesmus bijugatus isolated from soil. Plant and Soil 140: 25–34. Newman JR, Barrett PRF (1993) Control of Microcystis aeruginosa by decomposing barley straw. J. aquat. Plant mgmt 31: 203–206. Pillinger JM (1993) Algal control by barley straw: an interdisciplinary study. PhD Thesis, Open University, Milton Keynes, U.K.. Pillinger JM, Cooper JA, Ridge I (1994) Role of phenolic compounds in the antialgal activity of barley straw. J. chem. Ecol. 20: 1557–1569. Pillinger JM, Cooper JA, Ridge I, Barrett PRF (1992) Barley straw as an inhibitor of algal growth III: the role of fungal decomposition. J. appl. Phycol. 4: 353–355. Pillinger JM, Gilmour I, Ridge I (1995) Comparison of antialgal activity of brown-rotted and white-rotted wood and in situ analysis of lignin. J. chem. Ecol. 21: 1113–1125. Ridge I, Pillinger JM (1996) Towards understanding the nature of algal inhibitors from barley straw. Hydrobiologia 340: 301–305.

291 Ridge I, Pillinger JM, Walters J (1995) Alleviating the problems of excessive algal growth. In Harper DM, Ferguson AJD (eds), The Ecological Basis for River Management. J. Wiley & Sons, Chichester: 211–218. Tompkins J, Deville MM, Day JG, Turner MF (eds) (1995) Culture Collection of Algae and Protozoa, Catalogue of Strains. Natural Environmental Research Council, Ambleside, 204 pp. Van den Hoek C, Mann DG, Jahns HM (1995) Algae: An Introduction to Phycology. Cambridge University Press, Cambridge, 623 pp.

Webster J (1980) Introduction to Fungi, 2nd edn. Cambridge University Press, Cambridge, 669 pp. Webster JR, Benfield EF (1986) Vascular plant breakdown in freshwater ecosystems. Ann. Rev. Ecol. Syst. 17: 567–594. Welch IM, Barrett PRF, Gibson MT, Ridge I (1990) Barley straw as an inhibitor of algal growth I: studies in the Chesterfield canal. J. appl. Phycol. 2: 231–239.