Ecology and
biology of
nuisance algae
in rice fields
A report for the Rural Industries
Research and Development Corporation
by A.J. Grant, M. Pavlova, L. WilkinsonWhite, A. Haythornthwaite, I. Grant, D. Ko,
B. Sutton, & R. Hinde
School of Biological Sciences,
University of Sydney.
May 2006
RIRDC Publication No 06/010
RIRDC Project No US-115A
© 2006 Rural Industries Research and Development Corporation.
All rights reserved.
ISBN 1 74151 273 5
ISSN 1440-6845
Ecology and Biology of Nuisance Algae in Rice Fields
Publication No. 06/010
Project No. US-115A
The information contained in this publication is intended for general use to assist public knowledge and discussion
and to help improve the development of sustainable industries. The information should not be relied upon for the
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Research and Development Corporation, the authors or contributors do not assume liability of any kind
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Researcher Contact Details
A.J. Grant
School of Biological Sciences, A08,
University of Sydney, NSW, 2006.
Phone:
Fax:
Email:
02 9351 4488
02 9351 4119
[email protected]
In submitting this report, the researcher has agreed to RIRDC publishing this material in its edited form.
RIRDC Contact Details
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Level 2, 15 National Circuit,
Barton ACT 2600
PO Box 4776
KINGSTON ACT 2604
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[email protected].
http://www.rirdc.gov.au
Electronically Published in May 2006
ii
Foreword
Current methods of more intensive cropping and fertilising, together with flooding rice fields prior to
sowing pre-germinated rice seed, have increased competition for nutrients between rice and other
organisms. Competition from nuisance slime has been considered a factor in reducing the economic
potential of rice crops by some rice growers. Two forms of slime can reduce rice seedling
establishment. There are few data on this phenomenon with only one detailed study on algal slime,
done in 1980 (Noble & Happey-Wood, 1987). To maintain high productivity, it is important to
determine the source and identity of these forms of nuisance slime, and the conditions that favour the
growth of slime organisms over rice.
Green slime includes several species of filamentous green algae, single-celled green and blue-green
algae. We have now learned that farmers are readily able to manage green slime. However, farmers are
still concerned about brown slime which was previously believed to be caused by the growth of silicawalled algae, called diatoms (Noble and Happey-Wood, 1987). We have now determined that brown
slime is actually caused by bacteria, primarily iron-oxidizing bacteria. Bacteria produce a sticky slime
and many other organisms (invertebrates and green algae) and residue left from previous crops, bind to
it. Mats of brown slime up to 1 cm thick may form, covering small to large (2 x 50 m) areas, which
may hinder the emergence of rice seedlings during establishment.
This study revealed several conditions that lead to an increase in brown slime over rice seedling
growth. These conditions include high levels of urea combined with still water. We have suggested
changes in management practices for farmers that will reduce competition from brown slime.
This project was funded from industry revenue which is matched by funds provided by the Australian
Government through the Rural Industries Research and Development Corporation. It is an addition to
RIRDC’s diverse range of over 1500 research publications and forms part of our Rice R&D program,
which aims to improve the profitability and sustainability of the Australian rice industry..
Most of our publications are available for viewing, downloading or purchasing online through our
website:
• downloads at www.rirdc.gov.au/fullreports/index.html
• purchases at www.rirdc.gov.au/eshop
Peter O’Brien
Managing Director
Rural Industries Research and Development Corporation
iii
Acknowledgments
D. Patterson for identifying oxidized iron in the early brown slime samples, H. Hanert for identifying
species of bacteria, B. Singh for advice about biological iron oxidation, Malcolm Ricketts for help
with photography, Rice Research Committee delegates who were particularly helpful in providing
contact with farmers, DPI regional agronomists, S. Skinner of the Royal Botanic Gardens (Sydney),
scientists at CRC, Yanco, L. Parker of MIA, A. McLean of DIPNR, Leeton and the many farmers who
patiently advised us about their methods for rice growing.
Abbreviations
Within the results, individual farms are numbered and linked to the following codes for each locality.
Area code
Locality
CB
CL
DE
FN
GG
GR
HA
JE
LE
MD
ML
MU
TC
WA
WD
WL
Cobram
Colleambally
Deniliquin
Finley
Gogeldrie
Griffith
Hanwood
Jerilderie
Leeton
Morundah
Moulamein
Murrami
Tocumwal
Wakool
Widgelli
Willbriggie
Within this report, the following abbreviations are used.
Abbreviation
CRC
DPI
DIPNR
DLWC
MIA
MIL
PI
Full name
Co-operative Research Centre
Department of Primary Industries
Department of Infrastructure, Planning and
Natural Resources
Department of Land and Water Conservation
Murrumbidgee Irrigation Authority
Murray Irrigation Limited
Panicle Initiation (growth phase of rice)
iv
Contents
Foreword ............................................................................................................................................... iii
Acknowledgments................................................................................................................................. iv
Abbreviations........................................................................................................................................ iv
List of tables and figures...................................................................................................................... vi
Figures................................................................................................................................................ vi
Tables ................................................................................................................................................ vii
Executive Summary ........................................................................................................................... viii
Introduction ........................................................................................................................................... 1
1. Background ..................................................................................................................................... 1
2. Objectives of the project.................................................................................................................. 2
Methodology .......................................................................................................................................... 2
1. Information gathering...................................................................................................................... 2
2. Collection of soil and water samples and analysis of green slime .................................................. 3
3. Collection of soil and brown slime samples.................................................................................... 3
4. Microscopic examination of organisms in brown slime.................................................................. 3
5. Laboratory experiments determining the effect of flooding and urea on rice growth..................... 3
6. Consequences of brown slime for rice yield ................................................................................... 5
Results .................................................................................................................................................... 6
1. Information gathering...................................................................................................................... 6
2. Incidence of green slime in farm samples October 2002 ................................................................ 6
3. Quantity of green algae present in water supply channels and several farms ................................. 7
4. Brown slime collected from farms in October, 2002 ...................................................................... 9
5. Brown slime appears in several colours and forms ....................................................................... 10
6. Microscopic organisms found in brown slime samples................................................................. 13
7. Types of iron-oxidizing bacteria identified in samples of brown slime ....................................... 15
8. Brown slime and its management.................................................................................................. 16
9. Effect of sowing rice into damp soil versus flooded soil .............................................................. 20
9. Effects of fertiliser on the formation of brown slime .................................................................... 20
10. Selection of urease producing bacteria from brown slime samples ............................................ 22
11. Effects of fertilisers on rice seedling growth............................................................................... 23
12. Effect of pH on rice growth......................................................................................................... 25
13. Effect of brown slime on final crop............................................................................................. 27
Discussion............................................................................................................................................. 29
The nature of brown slime in rice bays ............................................................................................. 29
The link between urea and brown slime in flooded rice bays ........................................................... 30
Why is brown slime sometimes a problem?...................................................................................... 31
Conclusions .......................................................................................................................................... 33
Recommendations – suggestions for management of brown slime .................................................. 33
References ............................................................................................................................................ 36
Extensions ............................................................................................................................................ 38
Publications arising from this project ............................................................................................... 38
v
List of tables and figures
Figures
Fig. 1
Concentration of chlorophyll a in water samples taken from the Murrumbidgee River in
October 2002 (shaded bars) and August 2003 (stippled bars). Values represent the mean +
SEM (n=3). Error bars are absent from August 2003 (n=2).
Fig. 2
Concentration of chlorophyll a in water collected from rice farms and adjacent supply
channels in October 2002. Values represent the mean + SEM (n=3).
Fig. 3
Concentrations of chlorophyll a in water collected from water supply channels in June
2003.
Fig. 4A
Fig. 4B
Fig. 4C
Showing green algae growing in a mat of brown slime
A thin layer of brown slime that has remained at the soil surface.
An example of brown slime that has lifted off the soil surface but is not a problem but the
mat is sufficiently fragmented to enable rice plants to penetrate.
Showing rice seedlings having difficulty growing through a mat of brown slime
An example of organic matter in brown slime that has accumulated at the edge of a rice
bay
Fig. 4D
Fig. 4E
Fig. 5
Percentage of samples of brown slime from Table 4 in which iron deposits and
microscopic organisms were found (n = 30), collected from the soil (shaded bars, n = 14)
and water surface (open bars, n = 16), during November 2003 and November 2004.
Fig. 6
This photograph shows a diatom caught up in brown slime produced by iron oxidizing
bacteria collected from Site WD1 (photograph taken by D. J. Patterson). Bacteria are
barely visible at this magnification
Fig. 7
Siderobacter sp. bacteria and iron oxide present in a sample collected from site JE1 in
November, 2004.
Fig. 8
Photograph of bacteria predominantly Siderococcus sp., with iron oxide present in a slime
sample collected from JE1 in November, 2004
Fig. 9A
Growth of rice seeds in flooded (3 left-hand side tubes) versus damp soil (3 right-hand
side tubes) 10 days after sowing
Growth of rice seeds in flooded (3 left-hand side tubes) versus damp soil (3 right-hand
side tubes) 18 days after sowing
Fig. 9B
Fig. 10
Increase in iron oxidation with increasing urea (0, 58, 108, 198, 413 mg from left to right)
at 8 days after flooding
Fig. 11
Leaf lengths of the first three leaves to emerge from rice seeds.
Fig. 12
Showing a rice farmer measuring the pH of water in a rice bay, in November 2004.
Fig. 13
Showing brown slime during establishment (left-hand side) compared with the same crop
just before harvest (right-hand side).
Fig. 14
Three examples of grey soils that contain reduced iron (grey visible in footprints) beneath
a layer of oxidized iron (brown)
vi
Tables
Table 1
Summary of information gathering visits, October 2002–March 2005.
Table 2
Green nuisance algae identified in samples of green slime (n.d. = not determined).
Table 3
Samples of brown slime collected in October, 2002.
Table 4
Range of the numbers of microscopic organisms, other than bacteria, found in samples of
brown slime (n = 30) collected from the soil or water surface during rice establishment in
November 2003 and November 2004.
Table 5
Nitrogen source and amount and slime problems.
Table 6
Iron oxidation occurred in water above soil incubated with urea (810 mg) or urea plus
phosphate (54 mg) under flooded conditions in the laboratory for 7 days.
Table 7
Preliminary data showing that the addition of urea selects for soil bacteria that produce
urease.
Table 8
Range of pH in paddy water and soil samples measured between 23rd and 29th
November, 2004.
vii
Executive Summary
Introduction
This report describes an RIRDC-funded study of the causes and management of slimes affecting rice
production in the irrigated areas of southern New South Wales. At the start of the project it was
believed that two types, green slime and brown slime, were problematic and that both were caused by
algae – hence the title of our project. However, green slime is already very well managed and is,
therefore, a minor problem. Bacteria proved to be the major cause of significant brown slime
formation, and this became the major focus of our research.
The occurrence of slime in rice bays is important because it can reduce the yield of rice by killing
seedlings. Organisms in the slime also compete for nitrogen with the rice plants. Potential benefits of
the research include better yields of rice, lower fertiliser costs for farmers and possibly further
economies in the use of water.
Prior to this project, there had been very little research on the effects, causes and management of slime
in Australian rice production. The main work was that of Noble and Happey-Wood (1987).
Intended Audience
This report provides information on green and brown slime for rice farmers, agricultural extension
workers, educators and future researchers. It includes descriptions of the components and causes of
slime and recommendations for control, supported by field observations (by farmers themselves and
by researchers) and experimental evidence gained in our laboratory.
Background
Coherent, floating slime is common in rice fields. When rice seedlings are small, particularly before
the 3-leaf stage, slime may prevent them from growing through the water/slime surface layer, reducing
rice establishment. Even at later stages, slime may compete with the rice for nutrients.
Farming practices have changed in the last 20 years or so. Currently rice production is intensive, with
a crop each year and shorter periods of crop rotation; these two changes have led to more intensive use
of fertilisers, particularly urea. It is now more common to flood fields before sowing rice, rather than
after sowing into dry soil. Our research suggests that these changed practices have caused the increase
in nuisance slime reported by farmers.
Green slime is caused by algae – several types of filamentous green and blue-green algae form mats
that float on the water in rice bays. However, existing management practices deal with this effectively,
and farmers were not concerned about green slime affecting yields. Brown slime is stickier and more
tenacious, and may form mats up to 1 cm thick on the water surface; they may cover large areas (at
least 2m x 50m). These mats may reduce rice establishment, and are seen as a problem by some
farmers.
Previous work by Noble and Happey-Wood (1987) suggested that brown slime was caused by diatoms
– golden-brown algae with silica cell walls. We found that bacteria cause brown slime, and that
relatively few diatoms are found in it. Farmers variously attributed growth of slime to the presence of
decomposing organic matter, poor water circulation, poor land preparation and arrival of algae in
irrigation water. Evidence from their study led Noble & Happey-Wood (1987) to conclude that input
of algae from rivers was small and that conditions within rice bays increased algal populations. In
spite of this, farmers we spoke to were still concerned that algae from rivers or irrigation channels
might be causing slime. We were able to show that the algal populations of this incoming irrigation
water are, indeed, very low.
viii
Aims and Objectives
•
•
•
•
Identification and control of green slime
Determining the nature of brown slime, and why and when it is a problem.
Determining the conditions that increase brown slime
Suggesting strategies that farmers can use to reduce the brown slime problem
All objectives were achieved.
Methods Used
During the rice growing seasons of 2002 to 2004:
1. Information was gathered from rice growers in the western Murrumbidgee, Coleambally and
Murray Valley regions about the occurrence of nuisance slime and about which water and fertiliser
practices had changed over recent years. We collected information about irrigation supply water from
scientists in various relevant organizations.
2. Collection of soil and water samples and analysis of green slime: Samples of water and green
algal slime were collected from farms, from several sites on the Murrumbidgee River and from
irrigation supply channels (2002-3). Algae were identified and the chlorophyll content of the water
measured as a way to assess algal population size.
3. Collection of soil and brown slime samples: We visited farmers who had reported brown slime to
photograph the affected rice bays and to collect slime samples for microscopic examination. We also
collected soil samples from bays on a number of farms for laboratory experiments.
4. Microscopic examination of organisms in brown slime: We used these slime samples to identify
the main components of brown slime.
5. Laboratory experiments on the effect of flooding and urea on rice growth: Laboratory work
allowed us to use replicates and to test a number of different parameters rapidly and within our budget.
We did laboratory experiments to determine the effects of sowing into flooded soil vs. dry sowing and
of urea on rice growth.
Having found that bacteria, particularly iron-oxidizing bacteria, were the main cause of brown slime,
we also determined the effects of nutrients (particularly urea) and iron and the role of bacterial urease
(the enzyme that allows organisms to use urea) on slime formation in culture.
Farmers were concerned about the role of pH in slime formation, so we measured soil and water pH.
6. Consequences of brown slime for rice yield: Just before harvest, we revisited farms which had
experienced slime problems during seedling establishment in 2004 to find out if yield had been
reduced.
Results
1. Information gathering: Farmers and fellow scientists were
generous in providing information about the occurrence of slime, the
conditions believed to cause them and their effects on yields. These
data have been incorporated throughout the Results.
While green slime is
common, it is easily
controlled and does
not seem to affect
yield.
2. Collection of soil and water samples and analysis of green slime:
The green alga Spirogyra predominated in green slimes. The blueIt is caused by algae.
green alga Anabaena, and the green algae Oedogonium and
Hydrodictyon were sometimes present. Information from 11 farmers
showed that green slime was common but could be controlled well by lowering the water level or
adding a copper-based algicide (note that copper harms many organisms); one farmer used Round-Up.
Water samples from the river and the irrigation supply channels showed very low levels of chlorophyll
– that is, numbers of all algae (green and blue-green algae and diatoms) are very low in the incoming
water. The highest concentration measured was in a rice bay. We conclude (as did Noble & HappeyWood, 1987) that algal growth is encouraged, sometimes strongly, by conditions in the rice bays
themselves. No further study of green slime was warranted.
ix
3. Collection of soil and brown slime samples: Samples of brown slime from 6 farms contained
large populations of bacteria. They also contained diatoms, but too few to cause the slime or its colour.
We saw and photographed rice seedlings that were held back by brown slime. Control measures used
by farmers include dropping the water level, minimizing use of urea and recirculating or flushing the
water. One farmer noted that brown slime is not a problem if the rice is sod-sown. If water is kept
moving, either no film forms or it sticks to the soil, and does not affect the rice plants.
4. Microscopic examination of organisms in brown slime: Examination of further samples
confirmed the role of bacteria and showed that brown slime contains many types of organisms and
also organic debris, all stuck together by the “biofilm”
(biologically produced film) secreted by iron-oxidizing
Brown slime is caused by
bacteria. Without the bacteria to produce this sticky, coherent
bacteria. Algae are not
film, the other components of the slime would remain dispersed
important.
in the water.
The bacteria produce iron
oxide (rust) making the
slime brown.
Diatoms were present but rare - they were rarer than protozoa
and invertebrates. Green algae were insignificant. Other than
the large numbers of bacteria, the most interesting finding was
the presence of deposits of iron oxide in all but one sample of
brown slime. They give the slime its colour and their presence indicates that iron-oxidizing bacteria
are the main slime forming organisms.
5. Laboratory experiments on the effect of flooding and urea on rice growth:
Rice sown into damp soil (dry sown) grew faster than rice sown into flooded soil for the first 18 days.
Addition of urea before sowing into flooded soil inhibits the growth of the first 3 leaves of rice
seedlings, for the first 8 days after sowing. This is significant, since rice is vulnerable to being held
under by brown slime until the stronger third leaf is established.
Experiments in which bacteria were cultured in flooded soil collected from rice bays showed that (a)
no biofilm formed in controls (only water added) and oxidation of iron was slight; (b) addition of
phosphate alone did not cause the formation of a biofilm or oxidation of iron; (c) however, addition of
urea did cause the development of a biofilm and led to the oxidation of the soil iron, staining the
overlying water orange. Increasing the concentration of urea increased the rate at which iron was
oxidized.
Thus bacteria, iron and urea interact to form brown slime. Urease, an enzyme that breaks urea down, is
needed by bacteria to utilize urea. It is not produced by all
Brown slime needs urea,
bacteria. We showed that adding urea to soil samples increased
iron and iron-oxidizing
the amount of urease present, by selecting for soil bacteria that
bacteria to form.
produce it. No biofilm is formed unless urea is added to the
samples. If iron is added without urea, iron oxidation occurs
Iron and iron-oxidizing
but no film is formed. When both urea and iron are added a
strong biofilm is formed and iron is oxidized. Thus we
bacteria occur in all soils.
conclude that for brown slime formation, iron is required to
Urea levels are the key to
allow the growth of the iron-oxidizing bacteria that cause the
controlling brown slime.
film. However, without urea, their growth is too slow to form a
coherent film.
There was no correlation between pH and the presence of brown slime.
6. Consequences of brown slime for rice yield: We saw one site at which a patch of rice had been
lost due to persistent brown slime. However, the overall yield for the site was still considered good.
Implications for Stakeholders
Although some farmers were happy with the control measures they use for brown slime, others had
been less successful. We have shown that bacterial growth is the cause of brown slime and that urea
increases the growth of the relevant iron-oxidizing bacteria very strongly.
x
Evidence from our discussions with farmers and our observations show other factors that are involved
as well (see Recommendations, below), including the presence of still water during rice establishment.
Thus we have now provided a rationale for control measures for brown slime. This will benefit rice
growers and hence the whole industry.
Recommendations to rice growers
Suggested management practices to reduce brown slime.
ƒ
ƒ
ƒ
avoid using urea at rates above 57.5 kg N/ha in slime-prone soils prior to flooding
where possible, sow clover in rotation to fix nitrogen
dry sow rice or sow as soon as possible after flooding
If brown slime does form
ƒ keep water moving, especially in corners of bays, to prevent the formation of large slime mats
ƒ reduce water levels to keep rice leaves above slime BUT only after rice seedlings have
reached the 3 leaf stage
ƒ if the slime mats are limited, try raking the mats and throwing them onto nearby banks
How can farmers tell whether there will be a likelihood of brown slime
ƒ past experience of particular bays/paddocks
ƒ soil type; e.g. brown slime is more likely if the surface of grey soils always turns orange after
flooding, as iron is probably abundant. Many grey soils contain reduced iron.
ƒ about 3 weeks before sowing, test soil from bays thought to be likely candidates for brown
slime. Test by adding urea to soil plus water in a glass jar - watch for the appearance of
brown-orange coloured iron oxides in the next two weeks.
xi
Introduction
1. Background
In Australia, rice (Oryza sativa L.) is grown only in the irrigated regions of southern NSW
where about 2000 family farms produce about 1.2 million tonnes of rice a year. Rice
production in Australia is fully irrigated and 80-90 % of the rice crop is exported to more than
sixty destinations throughout the world where it competes against highly subsidised products
In the Murrumbidgee, Coleambally and Murray Valley regions, rice farmers have reported an
increasing problem of competition from both green and brown 'slime' which may hinder rice
seedling development. It has been assumed that these nuisance slimes are caused by the
growth of aquatic algae that may clog waterways as well as impeding the growth of young
rice plants, especially if conditions favour the growth of slime organisms over rice. Green
slime is caused by the growth of aquatic green algae and blue-green algae, which include
simple photosynthetic organisms that reproduce by spore production, or vegetatively by cell
division (Entwisle, 1998). Both filamentous and single-celled green and blue-green algae may
form green slime that clogs waterways and impedes the growth of young rice plants,
especially if conditions favour the growth of algae over rice. Brown slime has previously been
assumed to be caused by the growth of silica-walled diatoms (golden-brown algae) which
attach themselves to rice seedlings (Noble & Happey-Wood, 1987).
In 1980, a scientific study (Noble & Happey-Wood, 1987) attempted to estimate the
importance of the algal slime problem in rice fields of southern NSW. The response of 668
rice growers to a postal survey which included multiple choice questions asking about the
extent and incidence of algal slime problems, indicated that more than a third of the
respondents experienced a slime problem every 1-3 years. Some farmers believed that the
main factors linked to slime occurrence were the presence of large amounts of decomposing
organic matter, poor water circulation and poor land preparation (Noble & Happey-Wood,
1987). Other farmers claimed that they had an annual slime problem from algae that came in
with irrigation water. However, only small numbers of filamentous algae and diatoms were
found in the water coming from the Water Resources Commission channel. Once water was
retained in rice bays, the algal population increased both in quantity and species diversity.
These findings suggested that the water supply was not the prime source of algae and that the
conditions within the rice bays contributed to algal growth (Noble & Happey-Wood, 1987).
In their study, three sites (2 bays in each site) in the Denimein Irrigation District of the
Murray Valley were studied in detail over one season (Noble & Happey-Wood, 1987). Sites
included: one that had an annual problem of brown slime (considered to be due to diatoms); a
site that experienced a slime problem only rarely (type unspecified) and a site of natural
grassland that had not previously been used for rice. Prior to sowing and during the growth
period, algal species and biomass were determined in the incoming irrigation water and in rice
bays. Nutrients were also measured in soil samples (N, P, PO4, organic C, Cl, Ca, Mg, Na, K)
prior to irrigation and rice sowing. All sites were seeded by aerial sowing of Calrose rice.
However, no correlation was found between high levels of nuisance algae and either soil
nutrient levels or rice yield (Noble & Happey-Wood, 1987).
Rice farms range between 100 and 1000 hectares in size although only approx. 30% of each
farm may be sown to rice per year. However, the amount of rice sown each season is also
dependent upon the water allocation to each farm so that less rice may be grown during dry
years.
1
Current practices for growing rice vary considerably between individual farms but methods
include the following:
(i) drilling fertilisers into dry soil prior to flooding the bays with water. Nitrogen (N) and
phosphorus (P) are the main fertilisers added to rice crops (Ricecheck, 2003)
(ii) the main form of nitrogen added before planting has been urea because of its low cost and
ready availability. Recently, anhydrous NH3 has been used by some farmers which requires
specialised equipment for its application.
(iii) sowing rice into dry soil or sowing pregerminated rice seeds by air into flooded rice bays
(iv) adding herbicides into flooded rice bays and keeping the water 'locked up' (i.e. still) for
up to 10 days to control pre-emergent weeds
(v) flushing water through the rice bays to remove herbicides then adding permanent water
until just prior to harvest when the bays are drained.
(vi) another application of nitrogen may be made at panicle initiation (PI)
To reduce the quantity of brown slime in rice fields, Ricecheck (2003) has recommended
minimising decaying organic matter on the soil surface and lowering water levels to expose
both rice and slime to sunlight.
Nevertheless, the slime problem is now considered to be worse than 20 years ago which may
be due to changes in cultural practices. These changes include a preference for aerial sowing
into flooded bays rather than dry sowing, continuous rather than rotational rice crops and,
higher levels of residual soil nutrients due to more frequent fertiliser applications to sustain
continuous cropping. A continuing concern for some farmers is that increased levels of
nutrients enter rice paddies through the supply channel water. Although green slime has been
clearly linked to green algae, at the start of this project the cause and identity of nuisance
brown slime was still unknown.
2. Objectives of the project
•
•
•
•
Identification and control of green slime.
Determining the nature of brown slime, why and when it is a problem.
Determining the conditions that increase brown slime formation.
Suggesting strategies that farmers can use to reduce the brown slime problem.
Methodology
1. Information gathering
This project was conducted during the rice growing seasons of 2002-2004. We gathered
information from rice growers in the western Murrumbidgee, Coleambally and Murray Valley
regions about the location and seasonal appearance of nuisance slime and established which
water and fertiliser practices had changed over recent years.
We collected information about irrigation supply water from scientists at the Dept. of
Infrastructure Planning and Natural Resources at Leeton (DIPNR), and from the
Murrumbidgee Irrigation Authority (MIA) at Leeton and Murray Irrigation Limited (MIL) at
Deniliquin. We also contacted scientists at the Cooperative Research Centre (CRC) for
Sustainable Rice Production at Yanco Agricultural Institute and Charles Sturt University at
Wagga Wagga.
2
At field days, rice establishment meetings and during each field trip, we talked to farmers
who continued to have problems with brown slime and to those who successfully manage
brown slime.
2. Collection of soil and water samples and analysis of green
slime
During October 2002, we collected samples of green slime from 10 farms for identification of
algae. As several farmers still consider that the supply water to be the source of algal slime,
water samples (25-30 ml) were also collected from the Murrumbidgee River at Narrandera,
Gundagai and Jugiong in October 2002 and August 2003. Samples from several irrigation
supply channel sites collected during June 2003 were supplied by Dr. Mark Stevens of Yanco
Agricultural College for analysis.
We identified the predominant green algal species that were present (Entwisle et al., 1988) in
green slime samples using an Olympus, compound microscope. Green algae in water samples
were quantified using a Water pulse amplitude fluorometer which measures chlorophyll a .
3. Collection of soil and brown slime samples
Initially, we planned to study up to 10 rice farms and to collect water samples and soil
samples (core of 20 mm deep, 50 mm diameter) from rice paddocks and then examine several
parameters that cause algal slime. In the first field trip in October 2002, we collected samples
of both brown and green slime. After we found that green (algal) slime was manageable and
that brown slime was due to bacteria rather than to algae, we changed our approach and
concentrated on the problem of brown slime.
Farm selection for brown slime sample collection was made on the basis of
(i) farmers who contacted their local Rice Research Committee delegate and asked us to visit
them
(ii) farmers who contacted their local Department of Primary Industries (DPI) agronomists
with reports of nuisance brown slime
(iii) farmers who had heard of our project
Photographs of slime in rice bays were taken using either a Canon Ixus M-1 with Advanced
Photo System Fuji film or a Canon Powershot A70 digital camera.
4. Microscopic examination of organisms in brown slime
Samples of brown slime collected from farms were examined microscopically (Olympus CH2 compound microscope) and the types and numbers or organisms were counted in a 1.8 mm
field of view. Microscopic photographs were taken using a Zeiss Axiophot microscope
attached to a Leica DC 300F digital camera (Leica IM 1000 software, and OS98 operating
system) with the assistance of M. Ricketts.
5. Laboratory experiments determining the effect of flooding
and urea on rice growth
Variables such as climate, rain and wind are beyond the control of farmers. Thus we
concentrated on the parameters that are within their control.
3
To compare the rate of growth of rice seedlings in damp or flooded soil and the effect of
fertilisers, soil samples (up to 15 cm depth) collected from several rice farms were cultured in
sterile transparent 50 ml tubes (Greiner, Disposable Products, Australia). These tubes allowed
replicates of several treatments to be included in each experiment and being transparent,
enabled visual inspection of the soil surface. Tubes were placed in a growth cabinet with 13 h
light and a temperature range of 9 - 23oC based on the long-term average temperatures for
October (DPI, Deniliquin) to simulate conditions found in the rice growing region during rice
establishment.
5.1. The effect of flooding on early rice growth
To compare the effect of flooding versus sowing onto damp soil in early rice seedling growth,
urea was omitted. Soil (10 g) was added to each tube and distilled water added until the soil
was just damp or to a depth of 3 cm above the soil surface to simulate flooded conditions.
Water was added to compensate for evaporation during the experiment as necessary. Amaroo
rice seeds were soaked for 24 h in distilled water then drained for 24 h prior to dropping three
seeds into each tube
5.2. The effect of pre-sown nutrients on early rice growth
Yates superphosphate was used as a source of phosphorus in preliminary experiments. As
phosphate stimulated the growth of green algae but had no effect on the growth of brown
slime it was omitted from later experiments. Urea (Incitec) was included as the nitrogenous
fertiliser as it is the most commonly used form of nitrogen.
The addition of nitrogen is usually estimated from paddock history. For areas in which there
has been continuous rice cropping, RiceCheck (2003) suggests adding 180 - 240 kg N/ha but
lower amounts (0 - 60 kg N/ha) when subclover has been grown for the past 1-4 years. In 'cut'
areas where topsoil has been removed and not replaced, farmers are advised to add extra
nitrogen, up to 300 kg N/ha depending upon the depth of the cut (RiceCheck, 2003).
After visiting several farms and talking to farmers we realised that it would be impossible to
accurately simulate field conditions in test tubes. Urea is applied at the required number of kg
of N/ha, to the dry soil of rice paddocks over large areas by machinery. Nevertheless, within a
rice bay, due to a variety of spatial and physical parameters, the amount of urea will not
necessarily be uniform over each square centimeter of soil. To best simulate field conditions
in laboratory experiments we added urea at 27.5 mg per 50 ml tube as a rough approximation
of 180 kg N/ha.
To determine the effect of urea on early seedling growth under flooded conditions, urea was
added to the base of 50 ml tubes (in triplicate) at the following concentrations: none
(Controls), 27.5 mg or 55 mg. Ten g of soil (collected from rice paddocks) were then added to
each tube, followed by the addition of distilled water to a depth of 3 cm above the soil. The
tubes were placed in a growth cabinet with the lids removed (as described in Section 5). Rice
seeds were added to each tube three days after flooding.
5.3. Measurement of pH
The pH was monitored using Sigma pH strips or a TPS WP-81 pH meter.
4
5.4. Selection of urease producing bacteria from brown slime samples
Sterile tubes containing 25 ml of modified P medium (pH 6.1) for bacterial culture (KolbelBoelke et al., 1988) including Peptone (Amyl Media) as a source of nitrogen (Agudelo et al.,
2004), were inoculated with 100 μl of samples of brown slime supplemented with FeSO4 (700
μM) and urea (1 gm/L) as required. These cultures were further subcultured into fresh
medium at 4 week intervals. To measure urease activity, 12 ml of each culture were
centrifuged at 13,000 g for 5 min, the supernatant discarded and the pellets resuspended in
500 μl of 200 mM K2HPO4 buffer pH 7.6 in a 1.5 ml microfuge tube and frozen at -70oC for
24 h. After thawing, the pellets were sonicated (Bronson Sonifier B 12) for 3 rounds of 10
seconds each, keeping the tube on ice. Cell debris was pelleted by centrifugation and the
supernatant used to measure urease activity, using the NADH coupled assay at 25oC
(Kaltwasser & Schlegel, 1966). Non-specific NADH oxidase activity was determined in
samples run in parallel without urea and was deducted from the rates obtained in the presence
of urea. Amounts of urease were determined using a standard curve and Jack Bean urease
(Merck). Protein in the supernatant was determined in triplicate (20 μl aliquots) using the
fluorescamine assay (Udenfriend et. al., 1972).
6. Consequences of brown slime for rice yield
In March 2005, about 2 weeks before harvest, we revisited farms that had experienced brown
slime problems during the establishment period of November, 2004. We took photographs of
the same sites and talked to farmers to determine whether brown slime had reduced the
potential yield of the crop.
5
Results
1. Information gathering
Table 1 Summary of information gathering visits, October 2002 – March 2005.
sampling time
October 2002
August 2003
November 2003
March 2004
October 2004
November 2004
March 2005
August 2003,
2004, 2005
summary
Visited 10 farms, collected water and slime samples
Collected water samples from Murrumbidgee River (3 sites)
Information collected from DLWC and MIA, Leeton
Visited farms, collected 18 soil samples
Collected water samples from Murrumbidgee River (same 3 sites Oct ‘02)
Attended 8 rice establishment meetings, presented poster
Contact made with 55 farmers from Western Murray Valley and
Murrumbidgee regions
Collect 31 soil samples, 25 brown slime samples from 13 farms
Presented poster at rice information field day at Yanco Agric. Institute.
Spoke with scientists from Yanco, CSIRO Plant Industry, and soil
scientists at Charles Sturt University, Wagga Wagga.
Visited 13 farms.
Visited 11 farms, collected 27 soil samples
Spoke to 4 agric. Scientists at Yanco Agric. Institute
Visited 11 farms (same as in Oct ’04).
Responded to 7 other farmers experiencing brown slime problems
Presented talk to 130 farmers at 3 fields days at Yanco Agric. Institute.
Liaised with 3 scientists at Yanco Agric. Institute.
Visited 18 farmers.
Presented findings at annual RIRDC rice workshop at Yanco Agric.
Institute
2. Incidence of green slime in farm samples October 2002
The predominant species of green alga identified in green slime samples was Spirogyra.
Oedogonium, Hydrodictyon and Anaebaena (blue-green alga) were also present at 2 sites.
The methods of control were also noted (Table 2).
6
Table 2 Green nuisance algae identified in samples of green slime (n.d. = not determined).
*Coptrol is a commercial copper-based algicide.
Farmer
Code
Frequency
Perceived
source/cause
WD1
less common than
brown
annual, worse 1 in
3 years
water channel
CL1
a major problem 1
in 5 years
annual
organic matter,
nutrients
water channel
CL2
annual
water channel
JE1
occasional
GR4
GR5
occasional
n.d.
still weather
(dispersed by
strong wind
following day)
vegetative matter
n.d.
WL1
annual
water channel,
organic matter
applies CuSO4 or
*Coptrol
WL3
annual
organic matter
WL5
previous and
current season
n.d.
lowers water
level
n.d.
WD2
LE1
n.d.
Control methods main types
found in
sample
lowers water
Anabaena
level
lowers water
n.d.
level or adds
*Coptrol
lowers water
Spirogyra
level
lowers water
n.d.
level, drains, or
applies CuSO4
lowers water
Spirogyra
level
drains bays or
Spirogyra
applies CuSO4
applies Round-up
n.d.
Spirogyra
Spirogyra
Oedogonium,
Hydrodictyon
n.d.
Anabaena
Spirogyra
3. Quantity of green algae present in water supply channels
and several farms
The quantity of green microalgae, measured as chlorophyll a, was lowest in all water supply
channels and highest in a rice bay (CL1 RB 2, 92ng/ml, represented less than 1 ml of algae
dispersed in 100 litres of water).
Levels of chlorophyll a collected from 3 sites of the Murrumbidgee River were similar in both
October 2002 and in August 2003, at less than 8 ng/ml (Fig. 1). Chlorophyll a levels in water
samples collected from rice farms and adjacent supply channels in October 2002, also had
less than 20 ng/ml with one exception. In this case green algae were visible in this sample
collected from a rice bay at site CL1 RB 2 (Fig. 2).
Chlorophyll a levels in several water supply channels collected in June 2003 were also < 20
ng/ml (Fig. 3). Thus, there were fewer algae in the Murrumbidgee River than in water supply
channels.
7
Fig. 1 Concentration of chlorophyll a (in green algae) in water samples taken from the
Murrumbidgee River in October 2002 (shaded bars) and August 2003 (stippled bars). Values
for October 2002 represent the mean + SEM (n=3). Error bars are absent from August 2003
(n=2).
9
ng chlorophyll a/ml
8
7
6
5
4
3
2
1
0
Narrandera
Gundagai
Jugiong
Fig. 2 Concentration of chlorophyll a in water collected from rice farms and adjacent supply
channels in October 2002. Values represent the mean + SEM (n=3).
100
ng chlorophyll a/ml
90
80
70
60
50
40
30
20
10
0
WL5
BW
CL2
MC
CL1
MC1
CL1
MC2
CL1
RB1
CL1
FS
sampling site
Abbreviations of sampling sites for Fig. 2
Sampling site
WL5 BW
CL2 MC
CL1 MC1
CL1 MC2
CL1 RB1
CL1 FS
CL1 RB2
WD2 RB
WD2 FS
Farm code and source of sample
WL5 bore water
CL2 main channel
CL1 main channel mid-Oct
CL1 main channel late-Oct
CL1 rice bay 1
CL1 farm supply channel mid-Oct
CL1 rice bay 2
WD2 rice bay
WD2 farm supply channel
8
CL1
RB2
WD2 WD2
RB
FS
Fig. 3 Concentrations of chlorophyll a in water collected from water supply channels in June
2003.
18
ng chlorophyll a/ml
16
14
12
10
8
6
4
2
0
C1
C2
C3
C4
C5
R1
R2
W1
sampling site
Sampling site abbreviations for Fig. 3 are explained in the table below.
Sampling site
Location
C1
Main channel Yanco Agricultural Institute
C2
Channel opposite Yanco Agric High main gate
C3
Channel near bridge past Argyle St
C4
Channel near Leeton Dept. Agric. Field station
C5
Channel at Merungle Hill Rd. turnoff
R1
Murrumbidgee River, 10m upstream from Euroley Bridge
R2
Murrumbidgee River, Euroley Bridge
W1
Water wheel 796-3 near Whitton rice sheds
4. Brown slime collected from farms in October, 2002
Samples of brown slime were collected from 6 of the same farms in which green algal
samples were collected in October, 2002 (Table 3). Microscopic analysis of the slime
indicated that diatoms (golden algae) were present in all samples but in four out of six
samples, there was evidence of bacterial slime (Table 4).
Table 3 Samples of brown slime collected in October, 2002.
*Coptrol is a commercial copper-based algicide
Farmer
Frequency and Perceived
Control methods
Code
appearance
source/cause
WD1
main type of
algae found in
sample
lower water level,
circulate water
lowers water level
diatoms (and
bacteria)
diatoms
CL1
annual 'strangles' rotting organic
rice
matter
once in 5 years
increased organic
matter and nutrients
and climate
annual
hot and calm
lowers water level
CL2
annual
lowers water level
diatoms (and
bacteria)
diatoms (and
LE1
more prevalent in
some bays
9
Farmer
Code
Frequency and
appearance
Perceived
source/cause
Control methods
GR4
annual
high level of
vegetable matter
WL3
brown slime that
'pulls down rice
leaves'
dead/dirty green
algae plus silt
grazes sheep to reduce
vegetable matter and
lowers water level
(*Coptrol no effect)
lowers water level to
mud stage
main type of
algae found in
sample
bacteria)
diatoms
diatoms (and
bacteria)
5. Brown slime appears in several colours and forms
There is a range of colours found in brown slime, which consists of a mixture of different
organisms that adhere to the sticky slime produced by iron-oxidizing bacteria. It is the
bacterial slime that binds other components such as diatoms, organic matter and iron oxides,
giving it a firm structure.
Figure 4A shows a floating mat of brown slime that includes green algae that have become
attached to the slime produced by iron-oxidizing bacteria. In Fig. 4B, the layer of brown slime
has remained at the soil surface without hindering the growth of rice seedlings. In Fig. 4C,
while some brown slime has lifted off the soil surface and is now floating, it is in small
patches and has not hindered rice seedling growth. Figure 4D shows a thick (~1 cm deep)
layer of brown slime which is preventing the penetration of rice seedlings. Figure 4E shows
organic matter that has become attached to brown slime forming a mat which was blown to
the corner of a rice bay. In this instance some rice seedlings have become trapped in the mat.
In such cases, if the water is removed while rice seedlings are still at the 2-3 leaf stage, they
may be smothered and be unable to continue growing.
10
Fig. 4A Showing green algae growing beneath a mat of brown slime (area shown ~ 25 cm
across)
Fig. 4B A thin layer of brown slime that has remained at the soil surface (area shown approx.
30 cm across)
11
Fig. 4C An example of brown slime that has lifted off the soil surface but is not a problem
but the mat is sufficiently fragmented to enable rice plants to penetrate. Area shown approx.
30 cm across.
Fig. 4D Showing rice seedlings having difficulty growing through a mat of brown slime.
Area shown approx. 50 cm across.
12
Fig. 4E An example of organic matter in brown slime that has accumulated at the edge of a
rice bay. Area shown approx. 5 m across.
6. Microscopic organisms found in brown slime samples
Samples of brown slime were examined microscopically and the types of organisms noted.
Bacteria were present in all samples, but were not counted. Oxidized iron deposits were
present in all but one water sample. Diatoms were present in all soil samples and in most of
the water samples (Table 4) within a range of 0 to 49.
Table 4 Range of the numbers of microscopic organisms, other than bacteria, found in
samples of brown slime (n=30) collected from the soil or water surface during rice
establishment in November 2003 and November 2004. Field of view = 1.8 mm. Oxidized iron
deposits measured between 20 and 150 µm. Bacteria were also present in large numbers in all
samples.
Locality
Conargo
Deniliquin
Gogeldrie
Moulamein
Wakool
Willbriggie
Number
of
samples
2
2
3
7
6
10
Number of
oxidized
iron
deposits
3–3
1–3
2 – 14
0 – 33
1 – 20
2 – 35
Diatoms
(golden
algae)
Protists and
invertebrates
Green
microalgae
11 - 11
1-8
21 - 49
0 - 22
0 - 45
0 - 49
22 - 60
5 - 50
1 - 65
0 - 80
0 - 50
0 - 68
0-2
1-1
0-1
0-3
0 - 10
0-4
13
Fig. 5 Percentage of samples of brown slime from Table 4 in which iron deposits and
microscopic organisms were found (n = 30), collected from the soil (shaded bars, n = 14) and
water surface (open bars, n = 16), during November 2003 and November 2004.
% present in samples
100
80
60
40
20
0
Fe 3+
diatoms
protists
green
microalgae
Fig. 6 This photograph shows a diatom caught up in brown slime produced by iron oxidizing
bacteria collected from Site WD1 (photograph taken by D. J. Patterson). Bacteria are barely
visible at this magnification. Examples of a diatom, bacterial slime and oxidized iron are
indicated by arrows.
______________ Scale bar = 50 μm
14
7. Types of iron-oxidizing bacteria identified in samples of
brown slime
Dr. Hans Hanert, (Institute of Microbiology, Technical University of Brunswick, Germany)
who is an expert on the biology of iron oxidizing bacteria (Hanert, 1991a,b; 2002), identified
the following genera of bacteria present in brown slime samples; Leptothrix (possibly a new
species) Chlamydobacterium, Siderobacter and Siderococcus. Identification to species level
requires further analysis at the molecular genetics, electron microscopy and electron probe
levels. Because these analyses are both time-consuming and expensive, and identification was
peripheral to the aims of this study, they were not carried out. Figures 7 and 8 show examples
of Siderobacter sp. and Siderococcus sp. respectively.
Fig. 7 Siderobacter sp. bacteria and iron oxide present in a sample collected from site JE1 in
November, 2004.
Scale bar = 30 µm
15
Fig. 8 Photograph of bacteria predominantly Siderococcus sp., with iron oxide present in a
slime sample collected from JE1 in November, 2004.
Scale bar = 30 µm
8. Brown slime and its management
A number of farmers advised us that they have observed links between management practices
and the prevalence of brown slime. Table 5 lists information provided by 38 farmers and
shows the amount of nitrogen used (ordered from lowest to highest), the frequency of brown
slime and whether each farmer considered it to be a problem. The methods used by each
farmer to control brown are also included.
Table 5 indicates a link between increased amounts of urea and an increased incidence in
brown slime. Twenty-seven farmers had observed brown slime and 16 farmers reported it as
being a problem. The most commonly employed method of dealing with brown slime was to
manipulate water within rice bays. Eleven farmers dropped the water level and six
recirculated water. Several farmers mentioned a reluctance to drop the water level because of
the risk of losing water, a particular problem during drought. To minimise water loss and to
enable water to be recirculated one farmer at WA1 has recently build a ‘lake’ in which
drained water is collected for recirculation through rice bays.
Three farmers commented that waiting until the rice seedlings have grown beyond the 3 leaf
stage is important if brown slime is present. If the water is removed before the stronger third
leaf has appeared, the rice seedlings may be pulled down by the brown slime and remain
trapped in it at the soil surface. This point is illustrated in Fig. 4D
16
Table 5 Nitrogen source and amount and slime problems.
Site by code
Location
N as urea (kg
N as
N/ha) added
NH3 (kg
N/ha)
preflood
GR3
Griffith
14 - 28
WL4
Willbriggie
30 (fill area) to
172.5 (cut area)
JE2
Jerilderie
33
17
MU2
Murrami
35 - 57.5
GG7
Gogeldrie
37
WA1
DE5
Wakool
Deniliquin
40 - 83
46
MU3
GR2
Murrami
Griffith
FN1
GG3
DE2
FN2
GG6
LE1
ML2
Finley
Gogeldrie
Deniliquin
Finley
Gogeldrie
Leeton
Moulamein
85
57.5
57.5
64.4
72
72
72.2
84 - 113
Control measures
present in small area
slime in cut areas
not a problem
At >57.5 kg N/ha as urea sees
brown slime
56-112
57.5
brown slime
present but not a problem
a problem in some bays
present (more in black than in
red soils)
present, not a problem
not a problem unless >57.5 kg
N/ha as urea are added
sometimes a problem
not a problem
not determined
not a problem
sometimes a problem
not a problem
not a problem
when it appeared, he dropped
water after 3 leaf stage
avoids adding more than 57.5 kg
N/ha prior to sowing. If slime
appears,
drops water level but not until
rice is at 3 leaf stage
drops water level
recirculates water
recirculates water
avoids adding >57.5 kg N/ha as
urea
drops water until almost dry
zig-zags water
can always rescue if it occurs
N as urea (kg
N/ha) added
preflood
86
86 - 115
N as
NH3 (kg
N/ha)
brown slime
Control measures
present in bottom bays
a problem in some areas
not a problem
always present in some bays
flushes water through
18
Site by code
Location
ML1
MD1
MD2
TC1
Moulamein
Morundah
Morundah
Tocumwal
GG5
DE1
GG1
GG2
DE3
CB1
GR1
Gogeldrie
Deniliquin
Gogeldrie
Gogeldrie
Deniliquin
Cobram
Griffith
96
99
100
DE4
MU1
Deniliquin
Murrami
115
115
not a problem
occasionally if >57.5 kg N/ha
as urea added
present but not a problem
a problem for 5 days
JE1
WD1
Jerilderie
Widgelli
115
115
sometimes a problem
sometimes a problem
HA1
Hanwood
129
present sometimes
GG4
CL2
Gogeldrie
Coleambally
150
152
WL1
Willbriggie
161
not a problem
always a problem in some
bays
a problem
92
92 - 120
115
100
101
104
115
120
a problem even with NH3
present
present, not a problem
not a problem
not determined
drops water level
flushes with fresh water
but not a problem when sod sown
recirculates water
drops water level but not until
rice is at 3 leaf stage
drops water level
if water is stagnant for more than
1 week, recirculates water
drops water level until soil is just
damp
drops water level
Sprayed 12 kg/ha of CuSO4 at 2
leaf stage
Site by code
Location
WL2
CL1
WL3
Willbriggie
Coleambally
Willbriggie
N as urea (kg
N/ha) added
preflood
172.5
184
138 - 276
N as
NH3 (kg
N/ha)
brown slime
Control measures
a problem
a problem when calm and hot
often a problem, particularly
on grey soils
drains water
drops water level
19
9. Effect of sowing rice into damp soil versus flooded soil
Initial growth of rice sown onto damp soil without any addition of urea was faster than rice
grown into flooded soil, up until 10 days (Fig 9A). However, by 18 days, growth was similar
under both conditions (Fig 9B). Thus, at the crucial early stages of rice establishment up to 18
days, growth of rice grown under flooded conditions is relatively slow compared with rice
that is dry sown.
Fig. 9A Growth of rice seeds in flooded (3 left-hand side tubes) versus damp soil
(3 right-hand side tubes) 10 days after sowing
Fig. 9B Growth of rice seeds in flooded (3 left-hand side tubes) versus damp soil
(3 right-hand side tubes) 18 days after sowing
20
9. Effects of fertiliser on the formation of brown slime
In preliminary experiments we compared the effect of urea and phosphorus on brown slime
formation. Urea (810 mg) or phosphate (54 mg), or urea plus phosphate were added to the
base of 50 ml tubes and 10 g of soil were placed on top, then distilled water was added.
Visual symptoms of bacterial iron oxidation under laboratory conditions in this experiment
used an unrealistically high concentration of urea to ensure that the system was saturated.
After 7 days, a biofilm, which is the early stage of slime (O'Toole et al., 2000), had appeared
on the water surface of tubes containing either urea or urea plus phosphate but not in the tubes
containing phosphate alone.
Table 6. Iron oxidation occurred in water above soil incubated with urea (810 mg) or urea
plus phosphate (54 mg) under flooded conditions in the laboratory for 7 days. All distilled
water controls and phosphate only samples lacked the colour of iron oxidation. Colour ranged
from 0 (clear) to 3 (deepest shade of orange). The biofilm covered about 10% of the water
surface.
Farm
Soil
sample
Distilled
water
controls
Phosphate
Urea or
urea plus
phosphate
Biofilm which was
present in urea only
and in urea plus
phosphate tubes
CL2
FN2
FN2
GR2
LE1
MU2
WL3
1
1
2
1
2
1
2
0
0
0
0
0
0
0
0
0
0
0
0
0
0
2.5
<0.5
1
2
2.5
2
2
yes
absent
yes
yes
yes
yes
yes
21
Fig. 10 shows that the amount of iron oxidation is dependent upon the urea concentration.
Increase in iron oxidation with increasing urea (0, 58, 108, 198, 413 mg from left to right)
at 8 days after flooding.
10. Selection of urease producing bacteria from brown slime
samples
For urea to be used as a fertiliser it relies on the action of urease to release ammonia which is
further converted to ammonium ions which are then used by rice plants. This urease is
produced by soil organisms, typically by bacteria that are present in the 0 to 15 cm layer of
soil (Bremner & Krogmeier, 1989).
To further explore the link between urea and brown slime formation, we examined the
interactions between iron, urea and urease production. The results of preliminary experiments
are shown in Table 7. All tubes showed turbidity which is a sign of bacterial growth. It is
important to note that even in the absence of urea, nitrogen was supplied as Peptone
(hydrolysed casein) in the culture medium. Therefore, the results are not merely a response to
nitrogen in general.
Table 7 shows that in the absence of both iron and urea, the level of urease was low and a
white, readily dispersed biofilm was produced in the culture medium with a pH of 7.0.
Addition of urea alone led to an elevated production of urease and an increase in the pH of the
culture medium to 8.5 without biofilm formation.
The addition of iron alone did not elevate urease or pH but did lead to the production of the
characteristic colour of oxidized iron and the formation of a coherent biofilm. The
combination of iron and urea together led to an elevation of urease activity and pH, the
appearance of oxidized iron and the formation of a coherent brown biofilm (slime).
22
Table 7 Preliminary data showing that the addition of urea selects for soil bacteria that
produce urease. Details are described in Methods.
Dominant type of
bacteria
cocci + rods
rods + cocci
rods
rods with oxidized Fe
Appearance
(pH of culture
medium)
Selection medium
Urease
units/mg
protein
white, easily
dispersed
biofilm (7.0)
white
(8.5 - 9.0)
- Fe - urea
5.61
- Fe + urea
19.15
orange
(6.5)
coherent orange
biofilm
(8.5 - 9.0)
+ Fe - urea
6.87
+Fe + urea
16.41
11. Effects of fertilisers on rice seedling growth
The growth of the first leaf (Fig. 11a) was slightly inhibited by the addition or urea (27.5
mg/10 g of soil) between 5 and 23 days after sowing (tubes were flooded 3 days before
sowing). The second and third leaves did not appear in tubes containing urea until day 8.
Growth was lower in the presence of urea until day 15 after sowing when plants in urea (27.5
mg) treated soil exceeded growth of water controls (Fig 6b, c). That is, the presence of urea
depressed the early growth of rice seedlings. In the tubes containing 55 mg of urea, by 11
days, only 4 out of 9 seeds had produced one leaf (1-2 mm) and no further growth was
observed.
23
Fig. 11 Leaf lengths of the first three leaves to emerge from rice seeds. Leaf lengths of the
first three leaves to emerge from rice seeds (3 per tube) sown into water only (control, black
bars) and water with 27.5 mg urea (grey bars) three days after the addition of water to each
tube. Urea was added beneath the soil prior to the addition of water. Data represent the mean
(n=9) + SEM.
leaf length (mm)
(a) 200.0
leaf 1
150.0
100.0
50.0
0.0
5
6
8
12
15
19
23
days since sowing
leaf length (mm)
(b) 200.0
leaf 2
150.0
100.0
50.0
0.0
5
6
8
12
15
days since sowing
24
19
23
leaf length (mm)
(c) 250.0
leaf 3
200.0
150.0
100.0
50.0
0.0
5
6
8
12
15
19
23
days since sowing
12. Effect of pH on rice growth
Farmers are aware of the importance of soil and water pH in controlling a number of
processes important in the growing of rice crops, including the effect of high pH on
micronutrient supply. Farmers also raised a concern about a possible relationship between
high pH and brown slime formation. Therefore, at our third visit during rice establishment, we
encouraged farmers to measure the pH in rice bays themselves (Fig. 12) with the exception of
Farm MU3 in which pH was measured by a researcher. The data shown in Table 8 clearly
demonstrate that there was no relationship between high pH in the rice bay water and the
presence of brown slime. Equally there was no evidence to suggest that the pH varied
diurnally or as a function of air temperature (Table 8). Even though large mats were present in
two sites, brown slime was no longer a problem because rice leaves were tall enough to
remain above any brown slime.
25
Fig. 12 Showing a rice farmer measuring the pH of water in a rice bay, in November 2004.
The yellow coloured water is due to stirring up the brown slime present on the submerged soil
surface.
Table 8 Range of pH in paddy water and soil samples measured between 23rd and 29th
November, 2004. Rice seedling size (15-38 cm).
Farm
code
Site
within
each
farm
MU3
WA1
WA1
WA1
WA1
GG3
GG3
LE1
DE1
DE1
MU3
1
1
4
3
2
2
1
2
3
4
2
Brown slime
S = soil
surface
W =patches
on water
surface
M =large
mat on
water
surface
none
none
none
none
none
none
none
none
none
none
none
Water
6.45
6.90
6.98
7.02
7.05
7.15
7.40
7.50
7.62
7.67
7.80
pH
Top
soil
(5-10
mm
depth)
Deeper
soil
(~ 35 mm
depth)
Air
Temp
(oC)
Eastern
Summer
Time
6.18
6.43
6.70
6.91
6.82
6.98
6.78
7.84
6.48
7.35
6.66
5.96
6.06
6.18
6.03
6.31
6.80
6.48
6.70
5.98
6.22
6.16
37
28
28
28
28
34
34
37
34
34
37
1500
1700
1700
1700
1700
1200
1200
1630
1400
1430
1200
26
Farm
code
Site
within
each
farm
GG4
GG2
JE1
JE1
JE1
JE1
JE2
LE1
MU3
MU3
MU2
JE1
WL3
GG1
DE1
JE1
GG1
JE1
WA1
DE5
DE1
1
1
5
6
7
4
1
1
3
4
1
2
1
2
2
3
1
1
5
1
1
Brown slime
S = soil
surface
W =patches
on water
surface
M =large
mat on
water
surface
none
none
none
none
none
none
none
none
none
none
S
S
S
S
S
S
W
W
W
M
M
Water
7.80
8.62
8.63
8.68
8.72
8.76
8.84
8.85
8.94
9.70
7.52
7.67
8.50
8.58
8.79
9.00
7.94
8.00
8.50
8.08
8.72
pH
Top
soil
(5-10
mm
depth)
Deeper
soil
(~ 35 mm
depth)
Air
Temp
(oC)
Eastern
Summer
Time
7.45
8.62
7.65
6.26
6.55
7.42
8.85
7.16
8.20
9.40
7.05
6.60
8.00
7.90
7.20
8.90
7.70
7.15
6.78
7.00
7.02
6.80
7.15
6.55
6.15
6.17
6.81
6.82
6.78
n.d.
7.05
6.64
6.55
6.70
6.98
6.20
6.66
6.67
7.12
6.01
6.26
6.52
30
35
18
18
18
18
35
37
37
37
35
18
27
33
34
18
33
18
28
34
34
1000
1130
1200
1200
1200
1200
1230
1630
1400
1345
1140
1100
1800
1100
1330
1130
1030
1100
1730
1100
1220
13. Effect of brown slime on final crop
Figure 13 shows brown slime present in 3 sites at the time of rice establishment and again just
before harvest. There was no loss due to brown slime at WA1, in which the slime was
confined to small patches on the water surface. At DE5 Site 1, despite the thick layer of
brown slime that was present in November, the wind blew the slime out of the bay over the
bankless channel once the water level was raised. However, at DE5 Site 2, the brown slime
could not be shifted by water flow because of the uneven nature of the soil in this part of the
rice bay and rice did not become established there. Nevertheless, the farmer said that the loss
due to brown slime in this location had little effect on the overall yield of 10 tonne/ha.
27
Fig. 13 showing brown slime during establishment (left-hand side, A, C & E) compared with
the same crop just before harvest (right-hand side, B, D & F).
A
Farm WA1
B
ft
C
Site 1 at DE5
E
Site 2 at DE5
D
F
Each of these sites was photographed in November 2004 and again shortly before harvest in
March 2005. The area lacking rice plants in Site 2 at DE5 was attributed to being unable to
shift brown slime during establishment because of the uneven nature of the soil in this part of
the rice bay.
28
Discussion
After visiting ten rice farmers during our first field trip in October, 2002, it quickly became
apparent that nuisance green slime, caused by green algae and blue green algae, was an
intermittent but manageable problem. The predominant algae identified in samples of green
slime collected from rice bays were: Spirogyra, Anabaena, Oedogonium and Hydrodictyon
(Table 1). Farmers usually manage green slime by flushing water through the rice bays and in
severe cases, by using copper compounds (Table 1). However, it should be stressed that the
use of Cu2+ as an algicide should be avoided as it also causes oxidative damage to many other
organisms (Halliwell & Gutteridge, 1999, Grant et al., 2003).
Several farmers believed that the source of green slime was the supply water, due to nutrients
entering the Murrumbidgee River before it reached the Riverina district. At their request, we
sampled water from the Murrumbidgee River and several supply channels. We found that the
levels of green and blue-green algae were very low in water supply channels (Fig. 3) which is
in agreement with the previous study by Noble and Happey-Wood (1987). Their study found
that both algal abundance and species richness increased in flooded rice bays due to
favourable conditions prevailing within the rice bays. We then investigated the problem of
brown slime to identify the organisms responsible, to determine why and when it is a nuisance
and which conditions increase its prevalence. We were then in a position to suggest possible
methods to manage brown slime.
The nature of brown slime in rice bays
In the brown slime samples that were collected in the first field trip, we observed diatoms
(golden brown algae) which were considered to be the cause of brown slime by Noble &
Happey-Wood (1987), but also we consistently found bacterial slime and some orange lumps
that we did not recognise. For assistance with diatom identification, we approached D.J.
Patterson (Patterson & Burford, 2001) who identified the orange lumps as being oxidized iron
produced by iron-oxidizing bacteria. In the light of this observation, we then realised that we
would need to modify our original research plan.
Our approach to investigating brown slime was determined largely by current rice growing
practices in Australia. For example, because each rice farm may cover several hundred
hectares, comprehensive soil analysis of even ten sites would be time consuming, impractical
and well beyond the allotted budget for this project. Similarly, field trials would be possible
only for a limited number of sites and therefore would be impractical. Also, as a result of laser
guided landforming (used to improve water efficiency), the soil profile in rice bays is often
heterogeneous within quite small areas making it impossible to compare experimental plots
within the same bay. Therefore, we concentrated our efforts on talking to farmers to learn
how they managed brown slime and to determine any links between cultural and management
practices and brown slime occurrence.
We ascertained that brown slime contains a variety of organisms (Table 4). The slime is
produced by bacteria and the brown colour is due to oxidized iron produced by iron-oxidizing
bacteria. Virtually all bacteria produce polysaccharides on their membrane surfaces. These
polysaccharides (complex carbohydrates) may form a sticky biofilm which can lead to the
formation of slime. Because this slime is sticky many other organisms including invertebrates
and green algae will adhere to it. Organic matter left over from previous crops may also
become trapped in it leading to the formation of large mats. Leptothrix and Gallionella
bacteria are known to oxidize iron and it has been suggested that there is a wider range of
organisms not yet fully characterised, that are able to oxidize iron at neutral pH (Emerson &
Moyer 1997).
29
Dr. Hanert, an authority on iron-oxidizing bacteria, has tentatively identified several genera of
iron-oxidizing bacteria (Siderocapsa, Siderobacter, Chlamydobacterium) in brown slime
samples from Australian rice-growing areas. He has suggested also that a new species of
Leptothrix is present. Further studies are required to identify these bacteria to species level
and to study their physiology.
The link between urea and brown slime in flooded rice bays
Bacteria need water and nutrients to grow. Soil contains many bacteria but in a dry soil, they
may remain in a dormant state. If water only is added to soil, bacterial growth will be very
slow. However, once nitrogen is added, they will grow more rapidly and continue to grow
until the nitrogen is used up.
When urea is added to soil, ureases produced by both bacteria and fungi in the soil, hydrolyse
urea through several steps to ammonia and carbon dioxide. Any ammonium left in the soil
undergoes further reactions to produce nitrate which can be used by plants (Stitt, 2003).
Iron oxidizing bacteria that are present at the oxic-anoxic interfaces (~3 mm soil layer) use
Fe2+ as an electron donor. They also use CO2 as a source of carbon and NH4+ (ammonium) or
NO3- (nitrate) as a source of nitrogen. There are some species of iron oxidizing bacteria that
utilise anaerobic nitrate dependent iron oxidation at neutral pH (Benz et al., 1998). Banu et
al., (2004) showed a positive correlation between microbial biomass, total N and free iron in
NSW soils. Thus in the Fe2+ rich soils of rice growing regions in Australia (21 to 408 mg
useable iron/kg soil in the 0 to 10 cm layer, courtesy of H. Gill, Yanco Agricultural Institute),
flooded rice bays that have been fertilised with urea are ideal habitats for the growth of iron
oxidizing bacteria.
There are no reports of iron oxidizing bacteria producing urease and when growing them in
culture nitrogen is usually supplied as NH4Cl (Kucera & Wolfe, 1957). Therefore it seems
likely that in urea fertilised soils, iron oxidizing bacteria utilise nitrogen in the form of
ammonium that is released from urea by other urease producing bacteria. This sequence of
events would explain why there is a time lag between the flooding of urea fertilised soils and
iron oxidation. It would also explain why the amount of iron oxidation depends upon the
amount of urea present (Fig. 10). Our preliminary data (Table 7) show that the inclusion of
urea increases the growth of urease producing bacteria whether iron is present or absent.
However, in the presence of both iron and urea, a sticky orange-coloured coherent slime
containing oxidized iron is formed.
The preliminary data in Table 7 suggest that the formation of thick brown slime has a
specific requirement for urea over other nitrogenous sources. Thus, the growth rate of iron
oxidizing bacteria and the subsequent likelihood of brown slime forming, will be limited by
the amount of available nitrogen.
All ureases (plant, bacteria, fungi, invertebrates) are variants of the same Ni-dependent
enzyme and are found in the cell cytosol (Sirko & Brodzik, 2000). Urease hydrolyses urea to
NH3 and CO2 through several steps. The NH3 is incorporated into organic compounds mainly
by glutamine synthetase. Rice plants produce urease in both leaves and roots although urease
activity is three to five times higher in leaves (Gerendas et al., 1998). Thus, once rice roots
have become established, they will be able to utilise urea nitrogen. However, if rice seedlings
growing under flooded conditions are slow to become established, bacteria (and other soil
organisms that produce urease) will proliferate. Urease producers will release ammonium into
the water, thus providing nitrogen for the growth of iron oxidizing bacteria that form brown
slime.
30
Several farmers have noted a link between brown slime and higher amounts of added urea
(Table 5). In our experiments, we found that after the addition of water to soil containing
urea, the water above the soil surface turned orange which indicated oxidation of iron; the
depth of colour depended upon the concentration of urea (Fig 10). The colour change was
followed by the appearance of a biofilm in tubes that contained urea or urea plus phosphate,
but not those that contained phosphate alone (Table 5). Thus, oxidation of iron was dependent
upon the addition of urea. We rarely observed a complex mat of brown slime in laboratory
experiments but we deliberately omitted organic matter (which was present in field samples
of brown slime), from experimental tubes.
Some farmers believe that adding a large amount of urea is necessary to achieve a high yield.
However, a study by Kealey et al. (1994) found that applying urea at 120 kg N/ha before
aerial sowing of rice did not increase the yield significantly (P= 0.05) above that found with
60 kg N/ha.
We found that up to 15 days after sowing, rice leaf growth was faster in water controls than in
urea (Fig. 11). Equally, when urea of twice the concentration was used, growth of rice
seedlings was almost completely inhibited, suggesting that high levels of urea could be
harmful to early rice growth.
Toxic effects of urea on seed germination and seedling growth have been observed in several
crops and had previously been attributed to impurities in the urea itself or to a high pH
resulting from urea hydrolysis (Bremner & Krogmeier, 1989). However, a pH of 10.8 had no
effect on germination of wheat, barley, corn and rye seeds, and it was found that the
inhibitory effect was due to ammonia, produced by urease present in the 0 to 15 cm soil layer
(Bremner & Krogmeier, 1989). Addition of as little as 1 mg urea/g of soil inhibited seed
germination by 5 to 16% and inhibition increased with increasing urea concentration, up to
58% for 1.5 mg urea/g soil and 100% inhibition at 2.5 mg urea/g soil (Bremner & Krogmeier,
1989). That the release of ammonia was due to urease activity was confirmed as inhibition of
germination was prevented when the soil was autoclaved (destroying urease activity) or a
urease inhibitor was included (Bremner & Krogmeier, 1989). Thus, there is a clear
relationship between increasing urea, production of ammonia by urease activity and inhibitory
effects on early plant growth.
Urea nitrogen may be lost to the atmosphere as ammonia. Black et al. (1987), observed almost
twice as high a loss of urea-N by volatilization as NH3 when 300 kg N/ha was applied to the
soil surface compared with 100 kg N/ha.
These results suggest that as more urea is added, soil urease activity and ammonia production
increase resulting in much of the added fertiliser being wasted.
Why is brown slime sometimes a problem?
Brown slime usually forms on the soil surface of rice bays beginning as a biofilm and may
rise to the water surface and form a mat. If the mat is sufficiently dense it can strangle small
rice seedlings and prevent them from growing through the mat, thus reducing rice
establishment, which may in turn lead to a lower crop yield.
As there is still no accurate method available for measuring soil nitrogen (RiceCheck, 2003),
the amount of nitrogen that is added prior to rice sowing is estimated from previous crop
history. Following landforming, the amount of nitrogen added to 'cut' areas is higher, on the
basis that those areas contain less nitrogen after the removal of topsoil.
31
We also learned from the first visit that brown slime appeared about 2-3 weeks after sowing
soaked rice seed into flooded bays and was only a problem when rice was sown into flooded
bays. That is, it was not a problem when rice was dry sown.
Our results show that rice growth is slower in flooded conditions than when dry sown and that
when urea is added, growth is further slowed. Thus, when dry sown, rice has an advantage
over bacterial growth when competing for nutrients compared to flooded conditions.
There is a possible link between soil colour and the likelihood of brown slime formation.
Several farmers have noticed that brown slime is more of a problem in grey or black soils
(Table 5) than in red soil. Grey soils are often indicative of reduced iron and orange soils
indicate oxidized iron (B. Singh, personal communication). When farmers walked into
drained rice bays to examine rice seedling growth or to retrieve samples of brown slime, we
noticed that the layer of brown slime on the soil was quite shallow (Fig. 14). Beneath this
orange brown layer, the soil was grey suggesting that the soil contained reduced iron which
had been oxidized at the aerobic/anaerobic interface by iron oxidizing bacteria.
Many farmers recirculate the water within bays or lower the water level to control brown
slime (Table 5). However, several farmers commented that timing is critical when lowering
water during the rice establishment period, if loss of young rice plants is to be avoided. For
example, if the water is lowered when the rice seedlings are still at the second leaf stage, then
they are likely to be dragged down along with the brown slime. If farmers wait a few days
longer until the rice plants have produced their third (stronger) leaves, then they will be able
to stand above the slime when the water is lowered. Our laboratory experiments showed that
when sown into flooded conditions in the presence of urea (Fig. 11), the stronger third leaf
does not reach 5 cm until after day 15 and that although the second leaf may be 5 cm in length
by day 12, it is often curled over under the water (results not shown). Several farmers
commented that as part of their brown slime strategy, they lower the water level only after
rice seedlings reach the third leaf stage (Table 5).
There is much anecdotal evidence about brown slime and many statements are made about
rice growing which may not apply to all sites. That is because the rice growing areas include a
complex range of organisms and it is not only impossible to measure all the parameters
involved, but those parameters are constantly changing. Thus, within a given site there may
be considerable spatial and temporal variations. For example, during our conversations with
farmers, we were told that the pH in rice paddocks is known to vary diurnally: higher during
the day and lower at night. Several farmers considered that a high pH (> 8.0) is harmful to
rice seedlings and that a high pH misbalances micronutrients in the paddocks “as it ties up
zinc”. Therefore, during our third visit during rice establishment, we encouraged farmers to
measure the pH in rice bays themselves so that they could find out exactly what does occur.
There was no correlation between time of day (during daylight hours) and pH variation in the
rice bay water nor any correlation between high pH (>8.0) and the presence of brown slime
(Table 7). Our experiments suggest that there will not be a relationship between brown slime
formation and pH unless both urea and available iron are simultaneously present.
32
Conclusions
Rice sown into water grows more slowly than dry sown rice. Addition of urea prior to sowing
rice may further slow seedling growth under flooded conditions. Thus, when rice is sown into
flooded bays containing urea, there will be a greater chance for brown slime to grow faster
than rice and therefore, a likelihood of brown slime formation which may interfere with rice
establishment, and in some cases, may decrease the final yield.
Recommendations – suggestions for management of brown
slime
Iron oxidizing bacteria and iron cannot be removed from rice growing soils. Thus, the
conditions that favour their growth need to be managed to reduce competition for nutrients
between rice, bacteria and subsequent slime formation. However, there are clearly several
strategies that can be implemented to reduce the nuisance of brown slime, and several farmers
already apply these successfully.
Suggested management practices to reduce brown slime
ƒ
ƒ
ƒ
avoid using urea above rates of 57.5 kg N/ha in slime-prone soils prior to flooding
where possible, sow clover to fix nitrogen in rotation with rice crops
sow rice as soon as possible after flooding.
If brown slime does form
ƒ
ƒ
ƒ
keep water moving, especially in the corners of rice bays, to prevent the formation of
large slime mats
reduce water levels to keep rice leaves above slime BUT only after rice seedlings
have reached the 3 leaf stage
if the slime mats are limited, try raking them off and throw them onto nearby banks.
How farmers can tell whether there will be a likelihood of brown slime
ƒ
ƒ
ƒ
past experience of brown slime in particular bays/paddocks
soil type e.g. if the surface of grey soils always turns orange after flooding (Fig. 14)
test suspect soils. About 3 weeks before sowing, add some urea to a small soil sample
taken from bays where brown slime may be a concern. Place in a glass jar, add some
water, put in a warm place and watch for the appearance of brown-orange coloured
iron oxides (takes about 2 weeks).
33
Three examples of grey soils that contain reduced iron (visible in footprints) beneath a layer
of oxidized iron (brown).
Fig. 14 A Taken at MU1, Moulamein after draining the rice bay, November, 2003.
Fig. 14 B. Edge of rice bay at JE1, Jerilderie in November, 2004.
34
Fig. 14C. Corner of rice bay at TC1, Tocumwal, November, 2004.
35
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37
Extensions
Presentation at rice workshops.
RIRDC Rice Research Workshops at Yanco Agricultural Institute 2003, 2004 and
2005.
Direct Interaction with farmers:
Visited 40 farms as part of this study
Spoke at rice establishment meetings
November, 2003 presented our current findings at 8 rice establishment meetings in the
Western Murray Valley (Wakool and Deniliquin) and Murrumbidgee (Willbriggie and
Benerembah) regions (55 farmers)
And at Field Days:
Oral presentation of our work at the Rice Information Field Day at Yanco Agricultural
Institute, March 2004 (about 150 farmers).
Oral presentation at three field days held at selected farms, March 2005 (130 farmers)
Publications arising from this project
Grant, A & Wilkinson-White, L 2004, ‘Nuisance algae in rice fields’, Farmers’
Newsletter Rice R&D Edition, vol. 165 (summer), pp. 28.
Grant, A, Pavlova, M, Wilkinson-White, L, Grant, I & Ko, D 2005, ‘Ecology and
biology of nuisance algae in rice fields’, Farmers’ Newsletter Rice R&D Edition, vol.
171 (summer), in press.
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