Journal of Applied Phycology11: 285–291, 1999.© 1999Kluwer Academic Publishers. Printed in the Netherlands.

285

The relative sensitivity of algae to decomposing barley straw

Derek Martin∗ & Irene RidgeDepartment 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 bothlaboratory and field conditions. Bioassays were conducted on a range of algae to evaluate their relative sensitivitiesto straw-derived inhibitor(s). A range of sensitivities was found, including some species that were resistant tothe straw-derived inhibitor(s). A microcystin-producing strain ofMicrocystis aeruginosawas very susceptibleto decomposing barley straw. Bioassays usingEuglena gracilissuggest that the inhibitory compounds are notderived from the phototransformation of straw decomposition products and do not act primarily by inhibitingphotosynthesis. 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 prop-erties of barley straw (Hordeum vulgare) when de-composed in water in the field. Subsequent researchsuggested that the inhibitory component derived fromthe straw itself and not from the associated mycoflora,although the production of algal antibiotics by bacteriawas not ruled out (Pillinger et al., 1992). Pillinger etal. (1994) proposed the main algal-inhibiting factor(s)to be some form of oxidized polyphenolic compoundderived from lignin which was solubilized from thestraw. Laboratory studies suggest that the inhibitor(s)are algistatic, rather than algicidal (Gibson et al., 1990;Newman & Barrett, 1993). Barley straw is now inwidespread use as a method of algal control since itis relatively inexpensive and no adverse ecological ef-fects have been reported. The growth of macrophytesand animal life appear unaffected (Everall & Lees,1996) and barley straw has been used in potable wa-ter supplies, where no unusual amounts of the organicchemicals routinely monitored by the water industryhave been found (Barrett et al., 1996).

A number of other materials have also been foundto be anti-algal including brown-rotted wood (Pillingeret al., 1995; Ridge & Pillinger, 1996) and some leaflitters, in particular oak leaves (Quercus robur) (Ridgeet al., 1995). In the case of oak leaf litter, oxidizedtannins leached during the first stages of decomposi-tion may be the initial source of algal inhibition, butthe continued inhibition of algal growth over manymonths suggests that lignin degradation is the sourceof the inhibitor(s) in leaves and barley straw alike.

It has been suggested that diatoms appear to be res-istant to the inhibitor(s), since one species was foundto grow in tanks of decomposing barley straw (Ridgeet al., 1995). However, in field trials in a reservoir,Barrett et al. (1996) found that the planktonic diatomsAsterionella formosaand Tabellaria sp. were sup-pressed after the introduction of barley straw, althoughno truly comparable control was used in this study andcell counts had to be compared to those obtained fromthe reservoir in previous years. Desmids have alsobeen observed in ponds which had been treated withbarley straw or oak leaves, where the growth ofClado-

286

phora glomeratawas shown to be inhibited (authors’unpublished data).

In the laboratory Microcystis aeruginosawasshown to be inhibited with as little as 2.57 g dry massof barley straw m−3growth medium (Newman & Bar-rett, 1993) and Gibson et al. (1990) showed that anumber of green algae were inhibited, but with dosesof up to 104 g dry mass m−3 growth medium. In thefield, effective algal control can be achieved with aslittle as 3–5 g dry mass m−3 water although doses of20–50 g dry mass m−3 water provide a wider safetymargin (Ridge et al., 1995).

The evidence suggests that there are differences insusceptibility between taxa of algae, hence, this studywas undertaken to identify the relative sensitivity of arange of taxa to decomposing barley straw. Comparis-ons of the differing sensitivity/resistance of a range oftaxa could give clues as to the mode of action of theinhibitor(s) from the underlying physiology and mor-phology of the species in question. Test species werechosen because they were known to be able to growto nuisance proportions, or for their ability to toleratestresses such as high temperatures and desiccation, orwhere there were interesting structural features whichmay have affected sensitivity towards straw-derivedinhibitor(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. Asingle growth medium was not adopted, since me-dia were chosen to obtain the best possible growthof control cultures. The test species were grown inmineral media, apart fromEuglena graciliswhichwas grown in an organic medium that supported het-erotrophic nutrition. All stock cultures were main-tained by continuous sub-culture and both the stockand experimental growth media were buffered with20 mM 4-(2-hydroxyethyl) piperazine-1-ethane sulph-onic acid sodium salt. All of the stock cultures wereunialgal but no attempt was made to maintain themaxenically. The JM1 medium was based upon the JMmedium of Tompkins et al. (1995) modified by New-man & Barrett (1993) and buffered to pH 8.2. The WCmedium was after Guillard & Lorenzen (1972) andbuffered to pH 7.2. Both the DM and EG media wereafter 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; SAGstrains were obtained from Sammlung von Algenkul-turen, Göttingen, Germany and Sciento strains wereobtained from Sciento, Manchester, UK.Stichococ-cus bacillariswas obtained from the culture collectionof the Natural History Museum, UK.Tabellaria floc-culosawas isolated from Loch nam Brac, Scotland(58◦ 23′ N, 5◦ 6′ W) and the Loe Pool strain ofMi-crocystis aeruginosawas isolated from Loe Pool, aeutrophic lake in Cornwall, UK (50◦ 4′ N, 5◦ 17′W). The Loe Pool strain was assayed after only onesub-culture in the laboratory, while it still retainedits colonial form. The other strains ofM. aeruginosatested no longer retained their colonial form and grewas single cells. The AK1 strain ofMicrocystis aeru-ginosawas supplied by Prof. G. A. Codd (Universityof Dundee) and it produced microcystin in cultureat approximately 0.2µg microcystin-LR equivalentsmg−1 dry mass (G. A. Codd, pers. comm.).Nitzschiafiliformis var.confertawas isolated from a tank of oakleaves which had been decomposing in the light. TheSynechococcussp. was isolated from contaminatedgrowth medium, which had been accidentally heatedto approximately 35◦C.

Preparation of straw and bioassay

Active barley straw was obtained by incubating 2000 gdry mass m−3 in aged tap water, in darkness, at roomtemperature for three to six months with vigorousaeration (Pillinger et al., 1994). All bioassays wereperformed in 150-mL conical flasks. Wet straw wascut into pieces≤5 mm in length, placed into 50 mLof the relevant medium and then inoculated with 1 mLof an exponentially growing culture of the test species.The dry mass of straw was found by drying over silicagel, 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 fromabove by fluorescent tubes with continuous light of120 µmol m−2 s−1 PAR and shaken twice daily.Euglena graciliswas grown both in the dark (het-erotrophic growth) and the light (photoheterotrophicgrowth). The duration of the assays is shown inTable 1.

Biomass estimation

Wherever possible the yield of algae at the end of anassay was quantified by cell counting on a haemacyto-meter or Sedgewick-Rafter chamber (Table 1). Where

287

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

Phylum Species Origin Growth Biomass Duration

medium estimation of assay

method (days)

Bacillariophyta Asterionella formosa Sciento DM count 4

Tabellaria flocculosa Loch nam Brac DM count 5

Nitzschia filiformisvar.conferta decomposing oak X3 DM count 4

leaves

Chlorophyta Closterium ehrenbergii SAG 134.80 WC count 8

Staurastrum pingue SAG 5.94 WC count 7

Cosmarium biretrum SAG 44.86 WC count 7

Spirotaenia erythrocephala SAG 7.89 WC chla 8

Chlorella vulgaris CCAP 211/12 JM1 count 4

Scenedesmus subspicatus CCAP 276/20 JM1 count 4

Pediastrum boryanum Sciento JM1 count 4

Stichococcus bacillaris BM 83092907A JM1 count 5

Euglenophyta Euglena gracilis Sciento EG count 6

Cyanophyta Anabaena flos-aquae CCAP 1403/13B JM1 chla 8

Anabaena cylindrica Sciento JM1 chla 8

Aphanizomenon flos-aquae CCAP 1401/1 JM1 chla 8

Oscillatoria redekei CCAP 1459/29 JM1 chla 8

Oscillatoria animalis Sciento JM1 chla 8

Microcystis aeruginosa Loe Pool JM1 chla 4

CCAP 1450/6 JM1 count 4

Sciento JM1 count 4

AK1 JM1 count 4

Synechococcussp. contaminant JM1 chla 4

cell counting was made difficult by the size of cells,or where the algae were filamentous, chlorophyllaanalysis was carried out according to the method ofPillinger et al. (1994). Briefly, at the end of an ex-periment the contents of each flask where filteredthrough a glass fibre filter (GF/C Whatman) and thefilter plus residue were extracted with 100% methanol.Chlorophylla was quantified by measuring the absorb-ance at 665 nm minus a background turbidity readingat 750 nm on a SP6-550 UV/VIS spectrophotometer(Pye Unicam) on both raw extract and acidified ex-tract, to correct for phaeopigments (Marker et al.,1980).

Experiments were performed on a range of dosesof barley straw, using five replicate flasks for eachdose, and an analysis of variance was performed totest for significance. The results are expressed as the

mean dry mass of straw, of at least two replicateexperiments, required to obtain a 50% reduction inyield when compared to a control value. Although themass of straw which was employed in the bioassayscould be controlled, the exact amount of inhibitor(s)and/or its rate of release could not be controlled, sincethis depended on the microflora which developed onthe straw and the length of the decomposition period.For this reason the test species were categorized ac-cording to their sensitivities (Table 2). This allowedcomparisons to be made without relying on exact 50%inhibition of yield values since they were subject toslight 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 Microcystis aeruginosa

0–1000

Sciento 70

Loe pool 90

AK1 180

CCAP 2301450/6 230

Tabellaria flocculosa 295

Anabaena flos-aquae 375

Closterium ehrenbergii 450

Cosmarium biretrum 540

Asterionella formosa 667

Oscillatoria redekei 670

Spirotaenia erythrocephala 750

Synechococcussp 750

Staurastrum pingue 960

Susceptible Aphanizomenon flos-aquae 1370

1001–2000 Pediastrum boryanum 1500

Stichococcus bacillaris 1940

Slightly susceptible Euglena gracilis > 27001

> 2001 Chlorella vulgaris > 5400

Resistant Nitzschia filiformisvar.conferta –

(stimulation) Scenedesmus subspicatus –

Anabaena cylindrica –

Oscillatoria animalis –

1Result obtained fromEuglena gracilisgrown in the light

Results

The amount of straw required to obtain a 50% re-duction in yield for all of the species which wereassayed is shown in Table 2; in all cases inhibitionwas significant (p<0.001). Barley straw inhibitedthe two planktonic diatomsAsterionella formosaandTabellaria flocculosa, whereas the surface-associateddiatom Nitzschia filiformisvar. confertawas not in-hibited and growth was actually stimulated. Placodermdesmids (Cosmarium biretrum, Closterium ehrenber-gii andStaurastrum pingue) and a saccoderm desmid(Spirotaenia erythrocephala) were very susceptible tobarley straw. Members of the Chlorococcales werefound in all of the categories presented in Table 2 apart

from the very susceptible category. Cyanophyta werefound to be both susceptible and resistant, althoughfour strains ofMicrocystis aeruginosawere found tobe the most susceptible of all the species tested.

Growth of all the resistant species was stimulatedin the presence of decomposing barley straw. In allcases stimulation was at least 200% of the controlwith up to 4000 g dry mass barley straw m−3 medium(p< 0.001).

Euglena graciliswas only slightly susceptible tobarley straw, but susceptibility varied depending onwhether the cultures were grown in the light or thedark (Table 3). After three days growth the light-grown cultures were not inhibited whereas those thatwere grown in the dark were inhibited in the pres-

289

Table 3. Yield of Euglena gracilisgrown with decomposing barley straw(2727 g dry mass m−3 medium) indarkness and in light. Values aremeans±S.E. (n= 5). Values for lightand dark assays are compared to con-trols grown in the light and dark respect-ively (significant differences from thecontrol (100%) are shown as∗ p< 0.01,∗∗ p<0.001).

Time (days) Yield (% control)

Light Dark

3 96± 8 56± 6∗6 67± 3∗ 29± 8∗∗

ence of barley straw. After six days growth boththe light- and dark-grown cultures were inhibited, al-though the dark-grown cultures were inhibited morestrongly. The same pattern of inhibition was obtainedin three replicate experiments.

Discussion

Decomposing barley straw inhibited the growth ofplanktonic diatoms under the specified laboratory con-ditions. This is contrary to the report of Ridge et al.(1995) which suggested that diatoms as a group maynot be inhibited. The present study supports the fieldobservations of Barrett et al. (1996), who showed thatAsterionella formosaand Tabellaria sp. were inhib-ited when barley straw was introduced into a reservoir.N. filiformis was not inhibited, as was perhaps to beexpected, since it was isolated originally from a tankof decomposing oak leaves. Oak leaves have inhibit-ory effects comparable to those of decomposing barleystraw (Ridge et al., 1995).

Newman and Barrett (1993) suggested that Cyan-ophyta were particularly susceptible to decomposingbarley straw, but the results presented in Table 2 do notsupport this view. Although all of theMicrocystis aer-uginosastrains were the most susceptible tested, bothAnabaena cylindricaandOscillatoria animaliswereresistant. The AK1 strain ofM. aeruginosawas verysusceptible which, apparently, is the first time a knownmicrocystin-producing strain has been shown to besusceptible to barley straw in the laboratory or in thefield. The Loe Pool strain ofM. aeruginosa, which wastested while it still retained its colonial growth form,was also very susceptible. Thus, the mucilaginous

mass which makes up a colony ofM. aeruginosagivesno greater protection from straw-derived inhibitor(s)than that of single-celled forms ofM. aeruginosa.

The fact that inhibition ofEuglena gracilisoccursat all in the dark shows that the phototransformationof straw decomposition products into phytotoxic com-pounds, proposed by Barrett (1994), cannot accountfor the inhibition observed in laboratory assays. Thecurrent results withE. gracilissupport those of Cooperet al. (1997) who showed several species ofSapro-legnia to be inhibited by barley straw when grownin the dark. Saprolegniawas formerly classified inthe fungal class Oomycetes (Webster, 1980), but theircharacteristic heterokont zoospores suggest that theyare more closely related to ’algae’ (van den Hoek et al.,1995). Although the straw-derived inhibitor(s) appearnot to be acting on a photosynthetic process inE. gra-cilis, further work is required to determine whetherphotosynthesis can be discounted as the primary site ofaction for the straw-derived inhibitor(s) in other pho-totrophs. What is interesting is thatE. gracilisshows adelayed and weaker inhibition when it is grown in thelight rather than in the dark (Table 3).

The ability to grow photoheterotrophically mayincrease resistance to some toxins, as reported byMegharaj et al. (1992), who showed that two speciesof Chlorococcales were resistant to certain phenoliccompounds when growing photoheterotrophically, butnot when growing auto- or heterotrophically. Furtherexperiments using axenic cultures may be valuable todetermine more clearly the growth characteristics ofE. gracilis and to see if light confers some degree ofresistance over longer periods of time. However, in thecurrent assays the presence of decomposer organismswas a prerequisite for inhibitor release so, unless thestraw is chemically treated to allow abiotic inhibitorrelease (Pillinger, 1993), the use of axenic cultures isproblematical.

Although only one of the desmids tested wasknown to form dense populations (Staurastrumpingue), other desmids were tested to determinewhether differences in their cell wall structure (Brook,1981; Gerrath, 1993) could affect sensitivity to barleystraw. However, the results presented in Table 2 showthat all of the desmids were very susceptible to barleystraw.

The concentrations of barley straw required foralgal inhibition in the present laboratory study are lar-ger than those which are reported to be necessary inthe 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 bemore toxic under the dynamic conditions that occurin the field; a static test would underestimate tox-icity. Whether this is true with the proposed chemicalsreleased from barley straw requires further investiga-tion. The difference in dosage highlights the problemsof extrapolating these laboratory investigations to thefield. However, of the small number of species whichhave been found to be susceptible in field studies(in reservoirs),Asterionella formosa, Tabellariasp.(Barrett et al., 1996) andAphanizomenon flos-aquae(Everall & Lees, 1996) were all susceptible in thecurrent laboratory studies (Table 2).

All of the resistant of species were stimulated bydecomposing barley straw. Growth stimulation couldbe due to the presence of some unknown organic nu-trient or perhaps certain organic substances originatingfrom the barley straw are acting in a growth regulatorycapacity (Larson, 1978).

The inhibitory action is thought to be based onthe same principle in barley straw, leaf litter (Ridgeet al., 1995) and brown-rotted wood (Pillinger et al.,1995). The use of decaying plant litter (particularlybarley straw) to control nuisance algae may have im-portant implications from a water management pointof view, but of possibly greater significance is theimportance of these materials in natural ecosystems.Leaf litter and decaying wood are major inputs intofreshwater ecosystems (Webster & Benfield, 1986)and could influence the species composition and popu-lation growth of algal communities if conditions werefavourable.

The laboratory studies presented here and by Gib-son et al. (1990) suggest that the sensitivity of algaeto decomposing barley straw is not related to generaltaxonomic or structural characteristics; even membersof the same genus can differ widely in their sensitiv-ity. The fact that some species appear to be resistantto barley straw may have important implications forwater management, although no known nuisance spe-cies were shown to be resistant in this study. Furtherevidence is required to see whether these findings canbe extrapolated into the field where natural inputs ofplant litter may play a part in controlling algal biomassand/or species composition.

Acknowledgements

We thank Dr E. J. Cox for identification ofNitzschiafiliformis var. conferta, Dr B. A. Whitton for helpful

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

References

Barrett PRF (1994) Field and laboratory experiments on the effectsof barley straw on algae. In Hewitt HG (ed.), BCPC monograph59: Comparing field and glasshouse pesticide performance II.British Crop Protection Council, Farnham: 191–200.

Barrett PRF, Curnow J, Littlejohn JW (1996) The control of di-atom and cyanobacterial blooms in reservoirs using barley straw.Hydrobiologia 340: 307–311.

Brook AJ (1981) The Biology of Desmids. Botanical Monographs16. Blackwell, Oxford, 276 pp.

Cooper JA, Pillinger JM, Ridge I (1997) Barley straw inhibitsgrowth of some aquatic saprolegniaceous fungi. Aquaculture156: 157–163.

Everall NC, Lees DR (1996) The use of barley straw to controlgeneral 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 PhycologicalResearch 9. Biopress, Bristol: 79–192.

Geyer H, Scheunent I, Karte F (1985) The effects of organic envir-onmental chemicals on the growth of the algaScenedesmus sub-spicatus: a contribution to environmental biology. Chemosphere14: 1355–1369.

Gibson MT, Welch IM, Barrett PRF, Ridge I (1990) Barley straw asan inhibitor of algal growth II: laboratory studies. J. appl. Phycol.2: 241–248.

Guillard RRL, Lorenzen CJ (1972) Yellow-green algae with chloro-phyllide 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 colouredstream. Freshwat. Biol. 8: 91–104.

Marker AFH, Nusch EA, Rai H, Reimann B (1980) The measure-ment of photosynthetic pigments in freshwaters and standard-ization of methods: Conclusions and recommendations. Arch.Hydrobiol. Beih. Ergebn. Limnol. 14: 91–106.

Megharaj M, Pearson HW, Venkateswarlu K (1992) Effects of phen-olic compounds on growth and metabolic activities ofChlorellavulgarisandScenedesmus bijugatusisolated from soil. Plant andSoil 140: 25–34.

Newman JR, Barrett PRF (1993) Control ofMicrocystis aeruginosaby decomposing barley straw. J. aquat. Plant mgmt 31: 203–206.

Pillinger JM (1993) Algal control by barley straw: an interdis-ciplinary study. PhD Thesis, Open University, Milton Keynes,U.K..

Pillinger JM, Cooper JA, Ridge I (1994) Role of phenolic com-pounds in the antialgal activity of barley straw. J. chem. Ecol.20: 1557–1569.

Pillinger JM, Cooper JA, Ridge I, Barrett PRF (1992) Barleystraw as an inhibitor of algal growth III: the role of fungaldecomposition. J. appl. Phycol. 4: 353–355.

Pillinger JM, Gilmour I, Ridge I (1995) Comparison of antialgalactivity of brown-rotted and white-rotted wood andin situanalysis of lignin. J. chem. Ecol. 21: 1113–1125.

Ridge I, Pillinger JM (1996) Towards understanding the nature ofalgal inhibitors from barley straw. Hydrobiologia 340: 301–305.

291

Ridge I, Pillinger JM, Walters J (1995) Alleviating the problemsof 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) CultureCollection of Algae and Protozoa, Catalogue of Strains. NaturalEnvironmental Research Council, Ambleside, 204 pp.

Van den Hoek C, Mann DG, Jahns HM (1995) Algae: An Intro-duction to Phycology. Cambridge University Press, Cambridge,623 pp.

Webster J (1980) Introduction to Fungi, 2nd edn. CambridgeUniversity Press, Cambridge, 669 pp.

Webster JR, Benfield EF (1986) Vascular plant breakdown infreshwater ecosystems. Ann. Rev. Ecol. Syst. 17: 567–594.

Welch IM, Barrett PRF, Gibson MT, Ridge I (1990) Barley straw asan inhibitor of algal growth I: studies in the Chesterfield canal. J.appl. Phycol. 2: 231–239.