Download Genetic variability and nitrogenase activity of cyanobacterial communi-

Genetic variability and nitrogenase
activity of cyanobacterial communities associated with tropical seagrass
meadows (western Indian Ocean)
Mariam I. Hamisi
Stockholm University
©Mariam I. Hamisi, Stockholm 2010
ISBN 978-91-7447-001-7, pp 1-57.
Front cover: Intertidal seagrass bed exposed during low tide at Mjimwema
coast, Dar es Salaam, Tanzania. Photo: Linghua Xu.
Printed in Sweden by Universitetsservice,US-AB, Stockholm, 2010
Distributor: Department of Botany, Stockholm University
To my Late Father Mzee Issa H. Ndwata, a friend and the
best of my son (Babu Upara), whose sudden death just
few weeks before submission of this thesis shocked me!
And
To My Mama, Husband and Children!
Abstract
Tropical seagrass ecosystems are highly productive and important for sustaining marine life and associated coastal societies. In this study, the diversity and role of nitrogen-fixing cyanobacteria associated with five common
seagrass genera in coastal regions of the western Indian Ocean (WIO; Tanzania) were examined, as well as the impact of anthropogenic activities.
Cyanobacteria were characterized morphologically and genetically (16S
rRNA and nifH gene phylogeny), as were diel variations in nifH gene expression, NifH protein levels and nitrogenase activity. The results revealed
that WIO seagrass beds supported a rich cyanobacterial diversity and that
these represented approx. 83% of total clones obtained (DNA and RNA nifH
clone libraries). Non-heterocystous genera, such as Oscillatoria, Lyngbya,
Leptolyngbya, Phormidium and Microcoleus dominated, while heterocystous
morphotypes such as Calothrix were less frequent and unicellular morphotypes (e.g. Gloeocapsa, Chroococcus and Chroococcidiopsis) were few.
Additionally, the phylogenetic analysis revealed several novel uncharacterized cyanobacterial clades. Cyanobacterial composition and nitrogenase
activity varied over seasons and between the seagrass species. Day time nitrogenase activity originated primarily from heterocystous phylotypes, while
non-heterocystous filamentous phylotypes fixed nitrogen at night. The highest activity in the diel cycle was 358 ± 232 nmol C2H4 g-1 h-1 at 09.00 associated with epiphytes of the seagrass Cymodocea. Nitrogenase activity was
consistently lower in anthropogenically disturbed (eutrophication) seagrass
sites. Such data suggest that diazotrophic cyanobacteria may be a significant
source of ‘new’ nitrogen in the often oligotrophic coastal regions of tropical
oceans. It is also proposed that the rapid shifts in the cyanobacterial population and function found may also be used as early disturbance indicator in
coastal management practices.
List of Publications
This thesis is based on the following publications and manuscripts.
Papers will be referred to in the Thesis by their roman numerals.
I. Lyimo, T.J. & Hamisi M.I. 2008. Cyanobacteria occurrence
and nitrogen fixation rates in the seagrass meadows of the
East Coast of Zanzibar: comparisons of sites with and without
seaweed farms. Western Indian Ocean J. Mar. Sci. 7:45-55
II. Hamisi, M.I., Lyimo, T.J., Muruke, M. & Bergman, B. 2009.
Nitrogen fixation by epiphytic and epibenthic diazotrophs associated with seagrass meadows along Tanzanian coast,
Western Indian Ocean. Aquat. Microb. Ecol. 57:33–42
III. Hamisi, M.I., Díez, B., Lyimo, T.J., Ininbergs, K. & Bergman, B. - Cyanobacteria associated with the phyllosphere of
the seagrass Cymodocea rotundata: diversity, diel nifH expression and nitrogenase activity. (Manuscript)
IV. Hamisi, M.I., Mvungi, E., Lyimo, T.J., Mamboya, F., Österlund, K., Bergman, B., Björk, M., Díez, B. – Changes in the
epiphytic cyanobacterial community and diazotrophic activity on
the tropical seagrasses induced by varying nutrient regimes. (Ma-
nuscript)
My contributions to the Papers were as follows:
Planned and performed sampling, field and laboratory work, data analysis and writing of the Papers I-IV, except for the experimental set up,
biomass and photosynthesis determination for Paper IV, and part of the
phylogenetic analyses of Paper III and IV.
Contribution from supervisors and co-authors were incorporated.
Paper I and II are reproduced with permission from the publisher.
Additional relevant Paper not included in the thesis:
I. Hamisi, M.I., Lugomela, C., Lyimo, T.J., Díez, B. & Bergman, B. - Molecular and morphological characterization of
cyanobacteria in the alkaline-saline Lake Momela, Tanzania.
(Manuscript)
II. Mamboya, F., Lugomela, C., Mvungi E., Hamisi M.I., Kamukuru, A., & Lyimo, T.J. 2009. Seagrass–sea urchin interaction in shallow littoral zones of Dar es Salaam, Tanzania.
Aquat. Conserv. Mar. Freshw. Ecosyst. 19: S19–S26.
III. Lyimo, T.J., Mamboya, F., Hamisi, M., & Lugomela, C. Food preference of the sea-urchin Tripneustes gratilla (Echinodermata: Echinoidea) foraging on tropical coastal seagrass
habitats of Dar es Salaam, Tanzania. (Manuscript).
Table of Contents
Abstract ........................................................................................................... v
List of Publications ........................................................................................vi
Table of Contents ........................................................................................ viii
Abbreviations .................................................................................................. x
Introduction ................................................................................................... 11
Cyanobacteria ........................................................................................... 11
Background and taxonomy .................................................................. 11
Seagrasses................................................................................................. 13
Overview and distribution ................................................................... 13
Importance ........................................................................................... 14
Epiphytes and associated organisms .................................................... 15
Nitrogen in the seagrass ecosystem ..................................................... 15
Anthropogenic threats to seagrasses .................................................... 16
Biogeochemical cycles of nitrogen .......................................................... 16
Fixed nitrogen sources ......................................................................... 17
Biological nitrogen fixation ................................................................. 18
nif gene diversification ........................................................................ 20
Regulation of nitrogen fixation ............................................................ 20
Nitrogen fixation by marine cyanobacteria .............................................. 21
Open water ecosystems (pelagic) ........................................................ 21
Coastal water ecosystems (benthic) ..................................................... 22
Aim ............................................................................................................... 24
Comments on the methodology .................................................................... 25
Selection of the study sites ....................................................................... 25
Cyanobacterial characterization ............................................................... 28
Nitrogen fixation (acetylene reduction assay) .......................................... 30
Results and Discussion ................................................................................. 31
Cyanobacterial (field and morphological characterization) ..................... 31
Cyanobacteria genetic characterization .................................................... 34
Diazotrophs and nitrogenase activity ....................................................... 36
Anthropogenic effects .............................................................................. 40
Conclusion .................................................................................................... 42
Way forward ................................................................................................. 43
Acknowledgement ........................................................................................ 45
References ..................................................................................................... 47
Abbreviations
ARA
ATP
BNF
Chl a
Cr
Cs
DGGE
DNA
Hu
LM
N
NEM
P
PCR
PSII
Q-PCR
RFLP
RNA
RAPD
SD
SE
SEM
SSCP
Tc
TGGE
Th
WIO
acetylene reduction assay
adenosine triphosphate
biological nitrogen fixation
Chlorophyll a
Cymodocea rotundata
Cymodocea serrulata
denaturing gradient gel electrophoresis
deoxyribonucleic acid
Halodule uninervis
light microscopy
nitrogen
north east monsoon
phosphorus
polymerase chain reaction
photosystem II
quantitative polymerase chain reaction
restriction fragment length polymorphism
ribonucleic acid
randomly amplified polymorphic deoxyribonucleic acid
standard deviation
standard error
south east monsoon
single strand conformation polymorphism
Thalassodendron ciliatum
temperature gradient gel electrophoresis
Thalassia hemprichii
western Indian Ocean
Introduction
Cyanobacteria
Background and taxonomy
Cyanobacteria constitute a highly diverse prokaryotic group and inhabit a
wide variety of environments offering different climatic conditions, including
the most extreme ones, such as hot springs and deserts as well as cold areas in
the Antarctic (Whitton & Potts, 2000). Cyanobacteria evolved approximately
3.5 billion years ago (Schopf, 2000), and their ancestors introduced oxygenic
photosynthesis using water as electron donor. Through the evolutionary trend
cyanobacteria have remained a critical component of terrestrial and aquatic
ecosystems as primary producers, and are often important agents for nitrogen
fixation (Knoll, 2008).
Cyanobacteria are microscopic in size ranging from 1-80 μm in cell diameter
and show substantial morphological diversity. Following botanical nomenclature they were earlier classified as blue-green algae, based primarily on their
algal- and plant-like photosynthesis, as well as their distinct pigmentation
caused by phycobiliproteins (eg. phycocyanin and phycoerythrin) and chlorophyll a (Chl a). This classification depends on observations of field material,
represented by herbarium specimen (Rippka et al., 1979). Hence, the identification is based on phenotypic rather than genotypic characters, such as morphology of cells and filaments, shape of the terminal cells, presence or absence
of sheaths, gas vacuoles, motile hormogonia, nitrogen-fixing heterocysts and
resting akinetes/spores (Thuret, 1875; Gomont, 1892; Komárek & Anagnostidis 1998, 2005).
Following the discovery using electron microscopy in the 70´ties that the cell
wall is typically gram-negative and the subcellular arrangement typically prokaryotic (lack of membrane bound organelles), the group was reclassified as
cyanobacteria following the bacteriological approach of nomenclature (Stanier
et al., 1978). Identification based on physiological and genotypic characters of
live specimen in culture, cyanobacteria have been categorised into five principle morphological groups or sections, representing different ascending complexities (Rippka et al., 1979) as shown below:
11
I.
II.
III.
IV.
V.
Unicellular cyanobacteria that reproduce by binary fission or budding.
Unicellular cyanobacteria that reproduce by multiple fission.
Filamentous non-heterocystous cyanobacteria that divide in one plane.
Filamentous heterocystous cyanobacteria that divide in one plane.
Filamentous heterocystous cyanobacteria that divide in more than one
plane.
However, both the botanical and the bacterial approaches have drawbacks. For
example, the unavailability of appropriate media/conditions that can support
growth of all cyanobacteria still makes only a limited number of cyanobacteria
isolated from nature being culturable (Kumari et al., 2009). Also changes of
phenotypic characters due to environmental variations are common and were
considered as problematic using the culture independent approach (Anagnostidis & Komárek, 1989). Recently, the use of molecular techniques in the culture
independent approach was introduced to identify cyanobacteria in various habitats, but it may still be problematic to identify characters signifying individual
taxa (Komárek & Anagnostidis, 1998, 2005). To avoid these constrains, the
identification of cyanobacteria is currently often based on a combination of the
more traditional botanical and the more modern bacteriological approach.
Cyanobacteria should therefore ideally be classified using this combined set of
markers, using molecular data (as a genetic basis) combined with
structural/ultrastructural, phenotypic and ecological data, when ever possible
(Komárek, J. person. comm).
Several molecular and more recently genomic methods, such as whole genomic sequencing and metagenomic analyses, have in recent years started to be
used to establish the diversity of cyanobacteria using cultured species and specimens from natural communities. The molecular methods expedite the exploration of many habitats and have demonstrated that some cyanobacteria are
habitat specific, and many habitats contain a still largely undescribed genetic
diversity. The diversity may have been masked using simple morphological
features as a base for identifications (Garcia-Pichel, 2008).
Molecular methods currently being used include for example: DNA-DNA
hybridization, nucleotide (nucleic acid) cloning and sequencing, several polymerase chain reaction (PCR) based fingerprinting methods, like denaturing
gradient gel electrophoresis (DGGE), temperature gradient gel electrophoresis
(TGGE), restriction fragment length polymorphism (RFLP), randomly amplified polymorphic DNA (RAPD), single strand conformation polymorphism
(SSCP), DNA microarrays, as well as PCR independent methods like G+C
content.
Various marker genes, encoding house-keeping or functional genes, have
been useful to identify as well as to study the function of cyanobacteria. The
16S rRNA gene (partial and whole sequence) has so far been the most frequently and successfully used cyanobacterial identifier, and has expanded our
knowledge substantially when it comes to identification of cyanobacteria from
12
various natural habitats including tropical marine habitat (Nübel et al., 1997;
Lundgren et al., 2003; Uku et al., 2007; Díez et al., 2007; Bauer et al., 2008;
Foster et al., 2009b). The 16S rRNA gene is highly conserved between different species of Bacteria and Archaea and has become an important molecular
marker for microbial phylogenetic analysis (Woese & Fox 1977; Woese, 1987;
Woese et al., 1990; Case et al., 2007). The most commonly used 16S rRNA
cyanobacteria specific PCR primers set is that designed by Nübel and coworkers (1997). Comparison of 16S rRNA gene sequence analysis of natural
communities and strains from culture collections provides insights into differences (Garcia-Pichel et al., 2001) and/or similarities between cyanobacteria
taxa. However, the 16S rRNA gene is considered to be more conserved in
function and structure than protein encoding genes, and thus the genetic diversity found using 16S rRNA as marker may not sufficiently deeply reflect the
physiological diversity of the organisms present (Fox et al., 1992). In addition
to the use of the 16S rRNA gene, functional encoding genes such as the nifH
gene, encoding the small subunit of the nitrogenase enzyme complex (Zehr &
McReynolds, 1989; Omoregie et al., 2004), the hetR gene, encoding the key
protein for heterocyst differentiation (Janson et al., 1999; Vintila & ElShehawy, 2007), as well as genes involved in toxin biosynthesis (Jungblut &
Neilan, 2006; Jonasson et al., 2008) have been widely used in cyanobacterial
classification studies.
The nitrogenase encoding nifH gene has become a most extensively used
functional gene to genetically identify and differentiate among nitrogen-fixing
cyanobacteria (Zehr & McReynolds, 1989; Olson et al., 1998; Poly et al.,
2001; Falcon et al., 2004; Steunou et al., 2008). For instance, a recent study by
Steunou and co-workers (2008) represents a comprehensive analysis of cyanobacteria in a natural community (hot spring microbial mat), in which also the
diel expression of a large set of nitrogen-fixing genes (nifH, nifD and nifK)
were monitored using quantitative-PCR (Q-PCR). The use of the nif genes as
markers has indeed uncovered a great diversity of nitrogen-fixing cyanobacteria in for instance marine environments (Zehr & McReynolds, 1989; Church et
al., 2005; Foster et al., 2006; Bauer et al., 2008; Steunou et al., 2008), while
their significance as nitrogen-fixers still often remains to be determined.
Seagrasses
Overview and distribution
Seagrasses were once terrestrial angiosperms (flowering plants) that subsequently adapted to live submerged in shallow oceanic and estuarine waters
about 100 million years ago (Duarte & Gattuso, 2008). They spend their entire
life cycle in shallow seawaters, and preferably in oligotrophic waters. Seagrasses have reproductive organs (flower and seeds), roots, rhizomes, stems
13
and leaves as other plants (Björk et al., 2008). These features distinguish them
from macro algae, often sharing the same ecological niche. Seagrasses distribution is narrow, estimated to cover about 0.1-0.2% of the global Ocean area
(Duarte, 2002; Björk et al., 2008). About 60 seagrass species have been identified, which may often form prominent seagrass meadows composed of single
species or highly mixed stands (Short et al., 2007). Seagrass biomass and biodiversity increases towards the tropics (Fig. 1), particularly in the Indo pacific
region including East African coast, where most prominent seagrass stands are
formed (Short et al., 2007). So far, not less than 12 seagrass species have been
identified along the east African coast (Ochieng & Erftemeijer, 2003).
Figure 1: Global seagrass distribution and diversity. As seen in the map,
seagrasses are found in temperate as well as tropical regions. Note higher
diversity (darker green shades) close to the equator along the Indian and
Pacific oceans, including the western Indian Ocean, particularly in coastal regions of Kenya, Tanzania and Mozambique. (Source: Short et al.,
2007).
Importance
Despite their relatively low coverage and restricted diversity, seagrasses form
a valuable marine ecosystem component with a high productivity (Costanza et
al., 1997; Duarte & Chiscano, 1999; Orth et al., 2006). Its productivity is predicted to be as much as three folds higher than that of coral reefs and tropical
rain forests (Björk et al., 2008). They may constitute an excellent food source
for a variety of invertebrates and vertebrates, such as dugongs, turtles and various herbivory fishes, and be crucial breeding, nursery and feeding grounds for
migratory fishes from adjacent habitat like coral reefs and mangroves (Unsworth et al., 2008). Seagrasses also inhabited by several important commercial marine species, as such serves as a traditionally important fishing grounds.
14
In the western Indian Ocean (WIO) region the coastal societies receive most of
their protein from fishes in seagrass meadows (Torre-Castro & Rönnbäck,
2004). Additionally, seagrasses may also form a link between the more land
based mangrove forests and the more submerged coral reefs. Together they
form a most crucial ‘three-partnership’ in marine coastal ecosystems which
contribute to a healthy ocean. Seagrasses are important in this context via sediment trapping and stabilization through their extended root and rhizome systems. The often tightly packed and conspicuous seagrass ‘meadows’ may also
be used by a variety of marine organisms and serve various important functions. Thus seagrass communities are essential for the marine life in general
and also for the associated coastal societies.
Epiphytes and associated organisms
The high productivity of the seagrass ecosystem is based on the potent physiological activities of the photosynthetic seagrasses. These activities are also
most likely potentiated by associated photosynthetic and non-photosynthetic
microbes (prokaryotic and eukaryotic), as well as other epiphytic and benthic
organisms (Hemminga & Duarte, 2000). For instance, it has been estimated
that the epiphytic community may contribute up to 56% of the total production
(CO2 fixation) in seagrass beds (Morgan & Kitting, 1984; Moncreiff et al.,
1992). Epiphytic microalgae are known to be important food sources for the
herbivory inhabiting seagrass beds. The abundance of epiphytes also influencing the meiofauna abundance in seagrass meadows (Pinckney & Micheli,
1998; Yamamuro, 1999). Furthermore, epiphytic and epibenthic seagrass associated microorganisms suggested as potentially diazotrophic, which may play a
role in N-support and N-cycling within the seagrass ecosystems (Hamisi et al.,
2004; Uku et al., 2007).
Nitrogen in the seagrass ecosystem
Seagrasses are known to primarily flourish in oligotrophic waters (Vonk et
al., 2008), increased nutrient levels have been shown to negatively impact seagrass growth and productivity (Burkholder et al., 2007). On the other hand,
nitrogen is among the key limiting nutrient to sustain the high productivity of
the seagrass meadows (Vonk et al., 2008). Seagrass can utilize various dissolved inorganic and organic N sources in the water column and the sediments
(Welsh, 2000; Vonk et al., 2008). Seagrasses are therefore not restricted to
water column nutrients, but able to also exploit nutrients from sediment pore
water using their extensive root and rhizome systems (Hemminga, 1998; Björk
et al., 2008). Additionally, heterotrophic nitrogen fixing bacteria associated
with the seagrass rhizosphere predicted to contribute a bulk of nitrogen required in tropical and subtropical seagrass meadows (Patriquin & Knowles,
1972; Capone, 1988; Welsh, 2000). For instance, in the Gulf of Carpentaria
15
(Australia), biological N fixed is suggested to be incorporated into the seagrass
Zostera capricornia which support approx. 50% of N demand (O’Donohue et
al., 1991). However, in temperate seagrass ecosystems, the biological nitrogen
fixation contribute less to the seagrass productivity, for example, less that 12%
of the N used by Z. noltii in the Bassin d’Arcachon (France) originated from
the biological fixation (Welsh, 2000).
Anthropogenic threats to seagrasses
About half of the world’s seagrass meadows are threatened and the rate of
disappearance is approx. 110 km2 per year since 1980 (Waycott et al., 2009).
The coastal location of the seagrass ecosystems made them vulnerable to the
numerous ever increasing anthropogenic activities. These include sewage disposal, mariculture, boating, destructive fishing, construction works, sediment
alteration, dredging, pollution; and in recent decades, by the phenomenal increase in tourism (Short & Wyllie-Echeverria, 1996). Increasing coastal sediment loading and eutrophication are the most common negative anthropogenic
impacts affecting the productivity of the seagrasses. Sedimentation reduces
water clarity and light availability to the photosynthetic seagrasses, while increasing nutrient levels (eutrophication) stimulate the growth of the ephemeral
micro- and macrophytes, which in turn may out-compete the more slow growing seagrasses by covering their photosynthetic machinery.
The level of ecosystem destruction depends on the concentration and exposure times. At moderate exposures, seagrass shoot productivity may increase
(Uku & Björk., 2005), while at higher nutrient loads for prolonged periods
massive growth of epiphyte and epibenthic algae may severely reduce seagrass
activities (Björk et al., 2008). Eventually, this may lead to a total shift in the
seagrass community, which may also affect organisms associated with seagrasses.
Biogeochemical cycles of nitrogen
The availability of fixed nitrogen is a major factor limiting the primary production in plants, including seagrasses. Nitrogen is a fundamental element for
all organisms as it is a vital building block of all amino acids and proteins.
Nitrogen is the nutrient that together with carbon is building up the basic structure of the cells and thereby all organisms. Dinitrogen (N2) gas constitutes a
large reservoir ~78% of the Earth’s atmosphere. However, N2 gas is not readily
available for assimilation by most terrestrial and aquatic organisms, except for
a limited set of prokaryotes (diazotrophs). These have the ability to break the
triple bond of the dinitrogen molecule and transform it into a bio-available
form. To be available, the N2 molecule needs to be converted (fixed) to NH4+.
This process can be performed anthropogenically in the Haber-Bosch
16
processes to produce artificial nitrogen based fertilizers, a product that today is
globally spread in agricultural areas and in particular in the industrialized parts
of the world. Some nitrogen may also be fixed by natural lighting. However, of
the total nitrogen being fixed, the biologically based nitrogen fixation (BNF)
through diazotrophic bacteria constitutes the largest proportion of the ‘new’
nitrogen being administrated to the biosphere. The contribution by BNF is
today estimated to approx. 60% of the total, representing ~250 Tg of ‘new’
nitrogen being transferred into the biosphere (Galloway et al., 2004).
Fixed nitrogen sources
In the pre-human world, BNF was also the dominant source of N while today
the industrial N production (Haber Bosch process) has become a most important source of N for human food production (Galloway & Cowling, 2002). In
fact, the industrial fixation is predicted to exceed the natural BNF. Asia, Europe and North America account for nearly 90% of the increase in the artificial
nitrogen demand and use (Galloway et al., 2004) with a substantial increase
predicted for other regions of the world to come.
The production of artificial nitrogen fertilizers uses approx. 1–2% of the
world's annual energy supply (Smith, 2002), and recently the global production
was reported to be ~160 Tg N per year. This production is assumed to meet the
global N demand for food production to support increasing human population
(Smil, 2002). However a draw-back of this remedy is that, the anthropogenic
nitrogen sources result in accumulation of reactive nitrogen in the environment, as the input rate is higher than the nitrogen removal through denitrification (Galloway et al., 2003). These increases may have impacts on human and
ecosystem health such as loss in biodiversity and habitat destruction due to
eutrophication, now considered being a major threat to coastal waters (Waycott
et al., 2009). The oceans are highly susceptible to the N accumulation through
atmospheric deposition and land run off (Galloway et al., 2004), and may become major N sink. Negative consequences of N accumulation involved environmental eutrophication, global acidification and stratospheric ozone loss
(Galloway et al., 2003). The increasing availability of nitrogen over the earth’s
bioregions also alters the cycles of other elements such as carbon, phosphorus
and sulphur (Falkowski et al., 2000). The alteration may cause serious and long
term environmental consequences, including global climate changes (Gruber &
Galloway, 2008). For instance, increased nitrogen oxide and ammonia emission as a result of increasing reactive N in the environment tends to affect atmospheric carbon cycle and consequently climate. However, the magnitude is
unknown and need to be addressed.
BNF is still the most important source of N fixed in non-cultivated areas, including the 70% of the globe that is covered by water. Recent estimates suggest that BNF contributes about 110 and 140 Tg N per year from land and
ocean, respectively (Gruber & Galloway, 2008). BNF is of greatest signific17
ance in tropical regions (Capone et al, 2005), such as in countries in Africa and
Latin America (Galloway et al., 2004).
BNF is counteracted via denitrification by heterotrophic bacteria (2 NO3− +
10 e− + 12 H+ → N2 + 6 H2O) and the nitrogen fixed is again returned into the
atmosphere as dinitrogen gas (Gruber & Galloway, 2008). In the oceans this
conversion is also via anaerobic anammox (Anaerobic Ammonium Oxidation
NH4+ + NO2− → N2 + 2H2O), a process performed by planctomycetes bacteria
(Strous et al., 1999). In nature the BNF and denitrification/anammox is expected to favor a steady state condition (Codispoti et al., 2001). However, recently reviews suggest that human activity altered the global natural N cycle
(Gruber & Galloway, 2008). Increasing nutrient inputs is predicted in tropical
region (Galloway et al., 2008), which also calls for concern. Much of our
knowledge on nitrogen dynamics is based on data from the temperate part of
the world, despite the fact that tropical regions harbor a vast terrestrial and
aquatic biodiversity. Information on ecosystems responses and their translation
into effects on community structures, such as biodiversity changes or even
losses, in tropical regions are therefore urgently needed.
Biological nitrogen fixation
BNF is a complex and highly energy demanding process, encoded by approx.
20 nif genes, that reduces dinitrogen gas into ammonia, according to the following equation:
N2 + 8H+ + 8e- + 16 ATP
2NH3 + H2 + 16ADP + 16Pi
The reaction is catalyzed by the nitrogenase enzyme exclusively found in
some free-living and symbiotic prokaryotic organisms collectively known as
diazotrophs. The nitrogenase is a complex enzyme system that consists of two
major proteins components, namely, dinitrogenase (MoFe-protein) encoded by
nifDK genes and dinitrogenase reductase (Fe-protein) encoded by nifH. Both
components also contain molybdenum (Mo) (Fischer & Hennecke, 1984)
though in some species, Mo may be replaced by vanadium (vnfH) or iron
(anfH) (Eady, 1996).
The nitrogenase enzyme is rapidly and irreversibly inactivated by oxygen.
There is one exception, however, as nitrogenase in Streptomyces thermoautotrophicus is unaffected by oxygen (Ribbe et al., 1997). Diazotrophs evolved
several strategies to protect the nitrogenase from being inactivated by oxygen.
For instance, the heterotrophic bacteria (Azotobacteraceae) are unique in their
ability to employ an oxygen-labile nitrogenase under aerobic conditions which
is based on a high metabolic rate allowing oxygen reduction at the membrane
(Oelze, 2000). In oxygen evolving diazotrophs such as cyanobacteria, a variety
of strategies has evolved to cope with this incompatibility, primarily by sepa-
18
rating the two processes spatially or temporally or through a combination of
both (see Fig. 2).
A most efficient nitrogenase protection strategy is that built on a spatial separation, i.e. the development of specialized cells for nitrogen fixation (heterocysts) (Section IV and V), with thicker cell walls to reduce O2 penetration, and
lack of oxygen production via photosystem II (PSII) (Mulkidjanian et al.,
2008). All heterocystous cyanobacteria fix nitrogen. The development of the
protective heterocyst from a vegetative cell is a complex process with numerous genes being affected, the master gene being a protease encoded by the hetR
gene (Haselkorn, 2007). Unicellular (Section I and II) and most nonheterocystous cyanobacteria of Section III practice a temporal separation strategy. About 50% of them fix nitrogen and only do so during the nonphotosynthetic dark phase (night) (Bergman et al., 1997). However recently,
one unicellular cyanobacterium was found to lack PSII (Zehr et al., 2008). The
non-heterocystous cyanobacterium Plectonema fixes nitrogen under microaerobic conditions only (Section III), while Lyngbya fixes nitrogen during the
dark period of a light/dark cycle (Bergman et al., 1997). The non-heterocystous
genus Trichodesmium uses a combination of a spatial (diazocytes) and temporal separation strategy with nitrogen fixation taking place in the light period as
in heterocystous cyanobacteria (Berman-Frank et al., 2001; Lundgren et al.,
2005). The various strategies are illustrated in Figure 2.
Figure 2. Morphological and behavioral adaptations enabling nitrogen fixation in different cyanobacterial morphotypes of Section I-IV.
Gray shaded areas indicate darkness and the white areas indicate daytime. The double dashed-lines in the graphs designate nitrogen fixation
rates, while photosynthesis is symbolized by the black solid lines (reproduced from Berman-Frank et al., 2003).
19
nif gene diversification
The nitrogen fixation genes are distributed among eubacteria and methanogenic Archaea. Phylogenetic analysis of nifH and nifD has been a commonly
used approach to genetically characterize nitrogen-fixers. In these, the nitrogenase genes are divided into clusters denoted I-IV (Chien & Zinder, 1996; Zehr
et al., 2003). However, Raymond et al., (2004) proposed five clusters from the
analysis of nifH, nifD, nifK, nifE and nifN. The five clusters are largely consistent with the previously described four clusters, which are:
I.
II.
III.
IV.
This cluster consists of the ‘conventional’ Mo-containing nifH and
some vnfH.
This cluster is composed of the non-Mo, non-V containing anfH as
well as nitrogenase from some Archaea.
This cluster consists of nifH sequences from a diverse group of distantly related microorganisms many of which are strict anaerobes (e.g.
clostridia, sulphate reducers).
A divergent loosely coherent cluster of nif like sequences from Archaea and from the distantly related protochlorophyllide reductase
genes (involved in chlorophyll biosynthesis).
The sequencing of nif genes from a variety of organisms, specifically the nifH
gene, provided a large and still rapidly expanding database of sequences from
organisms representing a diverse array of terrestrial and aquatic environments.
The database for the nif genes has by now become one of the largest nonribosomal gene datasets of uncultivated microorganisms from natural environments. Different clusters of diazotrophs, with unique phylotypes, have been
found to inhabit different niches (Zehr et al., 2003). Phylogenetic analyses
based on the nifH gene sequence of diazotrophs, including cyanobacteria, is
proposed to correspond to the phylotypic groupings based on 16S rRNA gene
analyses (Zehr & Capone, 1996). It should however be noted that the presence
of nif genes does not always parallel with nif gene expression and nitrogen
fixation (Zani et al., 2000) and calls for some precaution.
Regulation of nitrogen fixation
Nitrogen fixation is regulated by several environmental factors that affect the
function of nitrogenase enzyme, of most important are oxygen and combined
nitrogen sources. Nitrogenase enzyme is oxygen sensitive and requires ATP
and an electron donor for activity. These two antagonistic needs of nitrogenase
enzymes impose physiological constraints on diazotrophs. Consequently, diazotrophs require regulation of nitrogen fixation (nif) genes in response to the
levels of fixed and combine nitrogen availability, as well as carbon, energy and
the external oxygen concentration.
For instance, nifHDK genes are expressed as one operon and are highly conserved between diazotrophs (Golden et al., 1991), and are regulated in re20
sponse to factors that control nitrogen fixation. Probably as a consequence of
the high energy demand it is important to fine tune the process to save energy.
Hence, the regulation of nitrogen fixation takes place in a hierarchical manner
at all organization levels (transcription, translation and activity level) and in a
complex and integrated fashion (Adams & Duggan, 1999; Haselkorn, 2007).
The regulatory networks may differ from species to species, and may respond
to multiple environmental parameters (Dixon & Kahn, 2004). For instance, the
regulation of nitrogen fixation genes in response to oxygen availability is controlled by four proteins the histidine kinase FixL, the histidine kinase RegB, the
anti-activator NifL and the σ54-dependent activator NifA (Dixon & Kahn,
2004). In symbiotic bacteria, FixL and NifA provide a hierarchical response to
the oxygen concentration while in other diazotrophs, NifA is not directly responsive to oxygen, but its activity is regulated by flavoprotein (NifL) that
senses the redox status. The nif genes respons to the combined nitrogen status
is regulated by the PII signal transduction proteins (glnB) that are covalently
modified under nitrogen limiting conditions. Many bacteria and cyanobacteria
contain more than one homologue of PII, enabling hierarchical regulation in
response to the level of fixed nitrogen (Forchhammer, 2004).
Nitrogen fixation by marine cyanobacteria
Diazotrophic cyanobacteria play double roles in marine primary production
by being able to fix the two key nutrients carbon and nitrogen, from the atmosphere (Kumari et al., 2009). The diazotrophs may be free-living in the water
column (pelagic) or attach to substrates (benthic) in shallow waters, or may
occur as epiphytes or function as endosymbionts in eukaryotic hosts, such as
diatoms and dinoflagellates (Carpenter et al., 1999; Foster & Zehr, 2006; Foster et al., 2009a). Hence, they occupy a diverse range of ecological niches
within the marine habitat, with the pelagic forms dominating quantitatively.
Open water ecosystems (pelagic)
In the oligotrophic surface waters of tropical and subtropical regions of the
world, the non-heterocystous and diazotrophic cyanobacterium of the genus
Trichodesmium is extremely abundant and contributes substantially to new
ocean nitrogen inputs (Lugomela et al., 2001, 2002; Lundgren et al., 2003,
2005; Capone et al., 2005; Capone, 2008; Foster et al., 2009a). This genus
alone is estimated to contribute up to 50% of the total biological nitrogen fixation in the open ocean. In contrast, heterocystous cyanobacteria such as Nodularia are important bloom formers in temperate waters such as in the brackish
Baltic Sea (Sivonen et al., 1989; Staal et al., 2007). The cyanobacterial-diatom
symbioses may host the only abundant heterocystous cyanobacterium in sub
tropical and tropical surface oceans, where they play a significant role as nitro21
gen-fixers (Bryceson, 1982; Villareal, 1991; Carpenter et al., 1999; Gómez et
al., 2005; Foster et al., 2006; Foster et al., 2009a). The cyanobacterial genus
Richelia is a symbiont in the diatoms Rhizosolenia and Hemiaulus hauckii
(Carpenter et al., 1999; Foster & Zehr, 2006; Foster et al., 2009a) and an epiphyte on Chaetoceros (Gómez et al., 2005; Foster & Zehr, 2006; Foster et al.,
2009a). A recent study also identified a variety of other novel marine cyanobacterial hosts (Foster et al., 2006). This indeed stresses the need for additional
screenings of symbiotic cyanobacteria and for diazotrophs in marine ecosystems. Furthermore, recent studies indicate that diazotrophic unicellular cyanobacteria may be more important actors in oceanic nitrogen fixation than hitherto assumed (Falcón et al., 2004; Zehr et al., 2008).
Coastal water ecosystems (benthic)
Benthic ecosystems may be considered as important sources of fixed nitrogen
for marine ecosystems as a whole. Indeed such coastal systems are grossly
under-studied. It is hypothesized that they may harbor an important microbial
diversity that needs to be further explored (Bauer et al., 2008). BNF may contributes to new nitrogen in benthic ecosystems such as in seagrass meadows in
which it was shown that about 50% of the nitrogen demand originated from
BNF by heterotrophic bacteria of the rhizosphere (O’Donohue et al., 1991). It
has also been suggested that epiphytic cyanobacteria may represent a major
source of fixed N to oligotrophic reef systems (France et al., 1998). Most of the
studies on benthic BNF in coastal ecosystems has primarily focussed on microbial mats (e.g. Stal et al., 1984; Díez et al., 2007; Bauer et al., 2008; Steunou et al., 2008), on coral reefs (Hewson et al., 2007; Charpy et al., 2010), on
sponges (Weisz et al., 2007; Mohamed et al., 2008), on mangrove forests
floors (Lugomela & Bergman 2002; Kyaruzi et al., 2003) and seagrass meadows (Capone & Taylor, 1980; Capone & Carpenter, 1982; O’Donohue et al.,
1991; Welsh et al., 1996, 2000). The nitrogen fixation is prformed by both
cyanobacteria and heterotrophic bacteria. So far however, most of the benthic
ecosystems studies in seagrass beds has focussed on bacteria as the nitrogen
fixers (Bagwel et al., 2002 & the references therein).
Recent morphological and molecular investigations showed that tropical benthic marine ecosystems, including seagrass meadows, also contain complex
assemblages of nitrogen-fixing cyanobacteria (Uku et al., 2007; Díez et al.,
2007; Bauer et al., 2008). It was therefore suggested that they may have been
underestimated as a source of new nitrogen. Cyanobacteria, being photoautotrophic in nature, were for instance found to be prevalent in the above-ground
parts (the phyllosphere) of common seagrasses in the western Indian Ocean
(Hamisi et al., 2004; Uku et al., 2007). They were encountered as epiphytes on
the seagrasses or were spread as biofilms on the surrounding sediments. Our
knowledge is still however limited in regards to their genetic diversity and
nitrogen fixation in tropical marine benthic ecosystems. Hence, their physiological functions and ecological significance still needs to be examined. In a
22
future global perspective, an evaluation and estimation of cyanobacterial (and
bacterial) based nitrogen budgets in coastal marine ecosystems are needed if
we are ever to construct accurate biogeochemical nitrogen budgets. This lack
of knowledge may be particularly serious for the productive Indian Ocean
coastal regions for which our knowledge is still rudimentary. Add to this the
negative effects of the ever increasing anthropogenic-based pressures on coastal ecosystems in general, both in terms of pollutants but in terms of affectors
disturbing the natural nitrogen fixation, which may be highly detrimental for
certain ecosystems.
23
Aim
This study aims at exploring the biodiversity, physiological function and ecological significance of tropical marine cyanobacteria associated with seagrasses, and analysing consequences of related anthropogenic activities. To
reach the aim the following specific objectives are undertaken:
1. To explore the occurrence of cyanobacteria in seagrasses ecosystem in
time and space (Paper I, III &IV)
2. To in detail characterize the identity of the epiphytic cyanobacterial
populations using a polyphasic approach (Paper III & IV)
3. To assess their nitrogenase activity and its regulation, including their
nif gene expression (Paper II, III & Paper IV)
4. To analyse the impact of anthropogenic activities and increased nutrient levels on the cyanobacterial diversity and their ecological functions
in the seagrass ecosystem (Paper I, II & IV)
An understanding of ecological links between the coastal ecosystems, such as
that inhabited by seagrasses, and the human societies being dependent on their
services, may demonstrate aspects of the life supporting capacities of these
ecosystems which are generally poorly known, and point to appropriate management precautions to be taken.
24
Comments on the methodology
Selection of the study sites
All study sites were located in coastal areas of the western Indian Ocean,
more specifically in waters along the Tanzanian coast, the sediments of which
are often covered by large seagrasses meadows (Fig. 1). The sites were selected based on the seagrass availability (species and abundance), and anthropogenic influences. In Paper I, the effects of seaweed farming on cyanobacterial occurrence and nitrogen fixation associated with seagrasses were investigated at two sites at the east coast of Zanzibar, Chwaka Bay and Jambiani (Fig.
3). For Paper II, the effect of sewage discharges on nitrogenase activity of epiphytic and epibenthic diazotrophs associated with the seagrass meadows were
determined at two sites: Mjimwema located at 06º 50' S, 39º 21' E and Ocean
Road at 06o 48.3' S, 39º 18' E (Fig. 3), both on the coast outside the city of Dar
es Salaam (Tanzania). OR is a more nutrient-rich site as it receives major inflows and the sewer pipeline that drains the Dar es Salaam city. The morphological and genetic characterization of epiphytic cyanobacteria, diel nifH gene
and protein expression, and nitrogenase activity were determined at the more
pristine site at Mjimwema in Paper III. For the mimicked nutrient enrichment
experiments in Paper IV, seagrasses were collected at the west coast of Unguja
(Zanzibar) Island and the experiment was performed in 20 L containers at the
Institute of Marine Sciences (affiliated to University of Dar es Salaam).
25
Figure 3. Map of coastal Tanzanian regions with the study sites indicated
by the number of the corresponding Paper in which these were examined.
Seagrass selection
Seagrasses belong to two families of the Monocotyledons, the Potamogetonaceae, and Hydrocharitaceae. There are 9 genera within Potamogetonaceae;
Zostera, Phyllospadix, Posidonia, Halodule, Cymodocea, Syringodium, Amphibolus, Heterozostera and Thalassodendron, while Hydrocharitaceae comprises the genera Halophila, Thalassia and Enhalus.
These flowering plants are prominent features along the east African coast
(Fig. 1). Based on the global seagrass bioregions the east African coast belongs
to the tropical Indo-Pacific region (Short et al., 2007). This is a vast tropical
region stretching from the east coast of Africa to the eastern Pacific Ocean and
holds more seagrass species than any other region in the world. Seagrass distribution in the tropical Indo-Pacific region can be found in a range of habitat
types from shallow estuarine environments to deep clear waters, but mostly
between the reef break and the shore, and at a depth of less than 10 m (Fig. 4).
26
Figure 4: Conceptual model for the seagrass distribution along the tropical
Indo Pacific region coastal waters according to habitat dominance. (After
Short et al., 2007).
The Indian Ocean along the east African coast hosts twelve seagrass species
within eight genera, namely: Halodule uninervis, Halodule wrightii, Cymodocea rotundata, Cymodocea serrulata, Syringodium isoetifolium, Zostera capensis, Halophila minor, Halophila stipulacea, Halophila ovalis, Thalassodendron ciliatum, Thalassia hemprichii and Enhalus acoroides (Ochieng &
Erftemeijer, 2003). Papers I-IV, focused on the intertidal seagrass meadows
(shallow waters) in which seagrasses are exposed during the low tides and
hence easily accessed. The intertidal zone is also assumed to be more exposed
to coastal anthropogenic activities. The five seagrass species, representing four
seagrass genera, used in this study were, H. uninervis, C. rotundata, C. serrulata, T. hemprichii and T. ciliatum. The gross morphology of these seagrasses
is presented in Figure 5. Selection was based on the fact that they are the most
common species in the intertidal zone along most of east African coast, including the study sites (Fig. 4). In Paper II, the four species H. uninervis, C. rotundata, T. hemprichii and T. ciliatum were used to obtain basic information on
the nitrogen fixation status of seagrasses in the east African region. In Paper III
and IV, C. rotundata and C. serrulata were used to in detail examine and characterize the cyanobacteria of the phyllosphere using a polyphasic approach
(genetic and morphological) and also their diel nifH gene and protein expression and nitrogenase activity. The selection of this particular genus for the
detailed characterization was based on the findings of Paper II and the previous
analyses performed by Uku and co-workers (2007). In Paper I, the seagrass T.
hemprichii was examined as it is common and abundant at the two study sites.
27
Figure 5: The morphology of the five studied seagrass genera. Magnifications of individual leaves are shown below the plants. Cr - Cymodocea rotundata, Cs - Cymodocea serrulata, Th - Thalassia hemprichii, Tc - Thalassodendron ciliatum, Hu Halodule uninervis. Reproduced from McKenzie et al.,
(2006-2010).
Some specific morphological features of the seagrass species examined are:
C. rotundata – leaves 2-4 mm wide with a rounded, smooth leaf tip and smooth
rhizome.
C. serulata - leaves 5-9 mm wide with serrated tip
T. hemprichii - hooked/curved shaped leaves with short black bars of tannin
cells in leaf blade, thick rhizome with scars between shoots.
T. ciliatum - cluster of ribbon-like curved leaves at the end of an erect stem,
round, serrated leaf tip and tough, woody rhizomes with scars from successive shoots.
H. uninervis – seagrass with narrow leaves less than 2 mm wide with trident
leaf tip and one central longitudinal vein.
Cyanobacterial characterization
Cyanobacteria were characterized using a polyphasic approach i.e. through
the use of both morphological (ocular observations and light microscopy analysis of the seagrasses and the leaves) and genetic tools (DGGE finger printing
analysis; cloning and sequencing of the epiphytic biofilms). The morphological
approach provides a rapid assessment of the diversity of the sample. However,
for the WIO region, this technique faced a challenge due to the lack of a regional specific identification keys for marine cyanobacteria. The only available
cyanobacterial key for the western Indian Ocean region is that of Silva & Pienaar, (2000). Although being comprehensive, it is restricted to marine cyanobacteria of the Kwa Zulu-Natal region (South Africa) and may not provide
enough coverage of the regional cyanobacteria flora further north. Relaying on
the commonly used identification keys, such as the one by Desikachary (1959)
and Komárek & Anagnostidis (1998, 2005) are not always fully appropriate for
28
specific cyanobacteria occurring in as seagrass epiphytes in specific tropical
marine waters.
The genetic characterization combining PCR and cloning or DGGE methodologies require good quality genomic nucleic acids in enough quantities to
ensure recovery of at least most of the cyanobacterial community present in the
sample investigated. For benthic marine environmental samples this is a challenge. There are numerous potential interfering substances and even inhibitors
in marine benthic samples associated with plants and sediments, such as salts,
plant debris, polysaccharides, polluting substances and the like. These influence the analyses from extraction to the amplification in the PCR reaction. The
use of different DNA and RNA extraction methods, including different commercial kits for DNA (GenElute Plant Genomic DNA Mini prep kit; SigmaAldrich Sweden AB, Sweden) and RNA (RNeasy Plant mini kit; Qiagen, German) may also per se generate some differences in the community composition
recovered. Extraction buffer quantities, along with other reagents, needed for
instance to be increased in relation to the amount of cyanobacterial/seagrass
sample used. Prior to the manufacturer procedures, a mechanical destruction
step was introduced using fast prep (FP 120, Thermo Electron Corporation,
USA) in order to increase the breakage efficiency of the cells and hence to
improve quality and quantity of the extracted nucleic acids.
Since cloning is a time-consuming methodological approach, the use of
DGGE for screening larger set of samples was the chosen approach in Paper III
and IV. Cloning was used in Paper III for additional comparative purposes and
in order to evaluate the data obtained from the DGGE analysis. However,
DGGE analyses may also include biases (Sekiguchi et al., 2001). The primer
selection, shown in Table 1 below, was based on our experiences from previous studies in another benthic system in the WIO region (Bauer et al., 2008).
Table 1: The 16S rRNA and nifH cyanobacterial primers used in Paper
III and IV.
Gene
16S rRNA
16S rRNA
16S rRNA
nifH
nifH
nifH
nifH
Primer
CYA106F
CYA781R(a)
CYA781R(b)
CNF
CNR
PolF
PolR
Sequence (5' to 3')
CGG ACG GGT GAG TAA CGC GTGA
GAC TAC TGG GGT ATC TAA TCC CATT
GAC TAC AGG GGT ATC TAA TCC CTTT
CGT AGG TTG CGA CCC TAA GGC TGA
GCA TAC ATC GCC ATC ATT TCA CC
TGC GAY CCS AAR GCB GAC TC
ATS GCC ATC ATY TCR CCG GA
Reference
Nübel et al., 1997
Nübel et al., 1997
Nübel et al., 1997
Olson et al., 1998
Olson et al., 1998
Poly et al., 2001
Poly et al., 2001
For DGGE primers CYA 106F and CNF had additional 40 nucleotide GC clamp at the
5´ end which is CGC CCG CCG CGC CCC GCG CCG GTC CCG CCG CCC CCG
CCC G
29
Nitrogen fixation (acetylene reduction assay)
The acetylene reduction assay coupled to gas chromatography (ARA-GC method) was the method of choice being inexpensive, sensitive and simple to use
under the ‘primitive’ field conditions given. As there are several experimental
factors that may affect the results the sampling strategies were set to minimize
these. For instance, removing the seagrass might affect the physiological performance of the seagrass and associated epiphyte. Seagrasses were removed as
gently as possible from the sediments, washing was avoided to retain attached
cyanobacterial epiphytes. Filtered sea water was added together with the seagrass (whole thalli or leaf pieces) samples to retain moisture inside the bottle
during incubation. The incubation bottles were also incubated in the original
place of the respective seagrass or in some experiments in a water container
filled with sediment and seagrasses in an open place to mimic natural environments. The controls were: (i), only acetylene, no seagrass (to identify ethylene contamination in the acetylene gas), (ii), no seagrass, no gas added (to
screen for a spontaneous production of ethylene from the seagrass/epiphytes)
and (iii) empty bottle (ethylene production from the bottle). These showed
negligible levels of ethylene produced in all experiments.
In most of the other studies seagrass nitrogen fixation analysis, intact cores
around the seagrass were used as previously described (O’Donohue et al.,
1991; Moriarty & O’Donohue, 1993; Welsh et al., 1996), in which the total
epibenthic and epiphytic diazotrophs of the seagrass in question is estimated.
Whilst, using the bottle approach, as in this study, the estimated nitrogenase
activity provides information on the specific contribution of either epiphytes or
epibenthos separately, hence making it easy to in detail study parts of interest.
No N15 measurement was performed to identify the true conversion factor between ethylene produced and nitrogen fixed. Rather the theoretical factor of
4:1 was used in Paper II, and whenever needed for comparative purposes. In
Paper I, III, and IV the nitrogenase activity was expressed as the amount of
ethylene produced. The reported mean acetylene to nitrogen conversion ratio
related to seagrasses is 3:1, the range being from 2.6 – 4.6, (Welsh, 2000).
However, to avoid over estimations the 4:1 ratio was used here.
30
Results and Discussion
Cyanobacterial (field and morphological characterization)
The seagrasses phyllosphere are apparently suitable habitat for cyanobacteria,
as they often accumulated and formed microbial biofilm on exposed surfaces,
as well as on adjacent sediments (Fig. 6). Structurally, cyanobacteria occurred
as dark/black ‘dots’ on the seagrass leaves, a behavior typical of heterocystous
cyanobacterial species, such as the genus Calothrix. The non-heterocystous
filamentous cyanobacterial genus Lyngbya was rather seen as short brown
(light to dark) restricted, or more spread, patches over the seagrass leaves.
However, the typical macroscopic Lyngbya majuscula aggregates occurred as
long entangled brown ‘filaments’ look like ‘hairs’ at the base of the seagrass
plants and adjacent sediments. Dark brown, olive green, reddish slimy biofilms, often also covering the adjacent sediments, were typically composed of
genera such as Oscillatoria, Microcoleus and also the thin filamentous nonheterocystous cyanobacteria of the Pseudanabaenacea family. The magnitude
of coverage varied from non-visible to thick microbial spread out biofilms
(Fig. 6), depending on e.g. the season as well as other natural and anthropogenically inflicted parameters affecting the shallow coastal waters.
Figure 6: Schematic illustration of the appearance and grade of
coverage of various types of cyanobacteria in the phyllosphere
of a seagrass. Note the gradual increase in cyanobacterial cover
(black areas) of the seagrass (exemplified by Cymodocea) from
left to right, and the differences in the cyanobacterial cover patterns, including on the adjacent sediments.
31
The cyanobacterial morphotypes varied considerably in size and shape as
evident under the light microscope. The cyanobacteria found represented distinctly different cyanobacterial morphotypes: ranging from unicellular and
filamentous non-heterocystous and heterocystous phenotypes. It was generally
found (Paper I, III and IV) that non-heterocystous filamentous cyanobacteria
dominated the cyanobacterial flora, followed by heterocystous filaments, and a
few intermixed unicellular morphotypes. Non-heterocystous cyanobacteria
found at all sites in Papers I, III and IV were Lyngbya spp. and Oscillatoria
spp. (Fig. 7 F-J). However, the thin non-heterocystous cyanobacteria of the
Pseudanabaenacea family (Fig. 7 K-N) predominantly concurred with the mentioned genera in the nutrient enrichment experiment in Paper IV, while the
bundle forming cyanobacteria Microcoleus sp. (Fig. 7 O) was also commonly
found in Jambiani (Paper I). The latter genus (Microcoleus) was rare in the
sites investigated in Paper III and IV. Calothrix spp. (Fig. 7 A-D) was the dominant heterocystous cyanobacterium at some stages in Paper III and IV, but
was not found in any of the sites in Paper II, while the heterocystous genus
Nodularia (Fig. 7 E) was seldom found. In general, heterocystous cyanobacteria have been rarely encountered in tropical open waters, but were recently
frequently detected in tropical benthic systems where they associated/attached
to seagrasses (Uku et al., 2007), were entangled in microbial mats (Bauer et al.,
2008) or appeared in relation to coral reefs (Charpy et al., 2010). Their presence in tropical benthic oligotrophic marine ecosystems, as repeatedly observed in this study, combined with their often highly efficient nitrogenase
activity suggest that they may play an important role in the nitrogen cycle of
these specific tropical ecosystems. The seagrass phyllosphere may provide a
stable ‘substrate’ for attachment and the tight often more calm seagrass stands
may promote the colonization of heterocystous cyanobacteria with fragile connections between heterocyst and vegetative cells (Stal, 1995).
Increasing human activities along the coast (Paper I) or moderately enhanced nutritional levels (Paper IV), led to a drastic change in the cyanobacterial population as well as a reduction in the nitrogenase activity. For instance,
these conditions promoted massive growth of microbial biofilm dominated by
larger (e.g. Lyngbya, Oscillatoria) and thinner non-heterocystous cyanobacteria (e.g. Leptolyngbya and Phormidium), while heterocystous species disappeared. Unicellular cyanobacteria of the genera Gloeocapsa, Chroococcus and
Chroococcidiopsis (Fig. 7 Q-S) were also identified and were commonly
mixed with non-heterocystous or heterocystous cyanobacterial epiphytes on
the seagrasses. The spiral coiled non-heterocystous cyanobacteria of the genus
Spirulina (P) was among our rarely encountered cyanobacteria taxa.
32
Figure 7: Micrographs of representative cyanobacteria associated with
seagrasses in tropical coastal areas of the western Indian Ocean. (A-E) the
heterocystous cyanobacteria known to be potential nitrogen fixers, (A-D)different Calothrix morphotypes, with A showing how they aggregate through
their terminal cells (heterocystous), B is a detailed filament from A; (E)- a
cyanobacterium tentatively identified as Nodularia; (F-J)- filamentous nonheterocystous morphotypes related to Oscillatoria and Lyngbya;(K-N) the thin
filamentous cyanobacteria (Pseudanabaenacea family). O- Microcoleus; PSpirulina; (Q-S) Unicellular cyanobacteria (Gloeocapsa, Chroococcus and
Chroococcidiopsis). Bars – 20 µm.
33
Cyanobacteria genetic characterization
To complement the morphological characterization, the first ever genetic characterization of cyanobacteria being epiphytic on seagrasses was conducted.
For the purpose the 16S rRNA and nifH genes were used as markers (Paper III
and VI). These analyses revealed the dominance among the microbes of a photoautotrophic cyanobacterial community in the light exposed seagrass phyllosphere. Previous studies had focused on bacterial diazotrophs of the rhizosphere
of seagrasses. Analysis using general bacteria nifH gene primers (Paper III)
showed that cyanobacteria represented approx. 83% of the total clones obtained from both DNA and RNA clone libraries.
Our genetic analyses (using cyanobacterial specific primers for 16S rRNA
and nifH genes; Table 1) recovered phylotypes representing all identified morphotypes. The sequences obtained in both Paper III and IV revealed a low percentage similarity, being <95% to known cyanobacterial sequences in the
GenBank, with the exception of a few 16S rRNA sequences related to uncultured phylotypes previously studied in the WIO or tropical regions (Lundgren
et al., 2003; Uku et al., 2007; Bauer et al., 2008). Lack of closely related sequences in the database was also shown in other Indian Ocean regional studies
(Bauer et al., 2008; Charpy et al., 2010). This clearly indicates the lack of genetic information in the benthic marine systems or the ocean as a whole in the
tropical Indian Ocean. Indeed, this region most probably contains some unique
cyanobacteria phylotypes, the identity of which needs to be further explored.
For instance, the phylogenetic analysis (Paper III and IV) illustrated the presence of new clades within the cyanobacterial radiation, which contains sequences only from seagrasses or from benthic related phylotypes, of tropical
marine ecosystems.
The phylogenetic reconstructions (16S rRNA and nifH genes) from Paper IV
contained similar sequences as those identified in Paper III, supporting the
accuracy in the phylogenetic affiliation of most sequences generated within our
studies. As seen in Paper III and IV, the 16S rRNA phylogenetic affiliations
verified that most of the recovered sequences were affiliated to the nonheterocystous phylotypes, previously described in different marine environments. For instance, several of the sequences obtained were closely related to
planktonic Oscillatoriales of the genus Trichodesmium, and the benthic genera
Blennothrix or Hydrocoleum. However, many of the sequences obtained
formed separated clades only related to unidentified seagrass associated cyanobacteria, which are closely related to sequences retrieved from work in this
thesis only (Paper III and IV). Similarly, sequences distantly related to Pseudanabaenacea family members formed clades of their own, only closely related to
other uncultured or unidentified phylotypes previously reported (Uku et al.,
2007; Bauer et al., 2008). These clades may therefore represent members of
hitherto unidentified or novel groups of cyanobacteria.
34
Figure 8: Part of phylogenetic tree of the partial (359 bp) nifH gene sequences of representative epiphytic cyanobacteria recovered from Paper III
and IV. The sequences were generated via nifH based DGGE and clones. The
tree was constructed from distance approximations by the NJ method and Kimura two-parameter in PAUP (version 4.0b10). The sequence of Methanocaldococcus jannaschii (L77117) was used as out group (not shown as the tree
35
represent only cyanobacteria part). The bold sequences in the tree represent
sequences from this study (Paper III & IV). The non bold seagrass associated
sequences in the tree are our own unpublished sequences from the seagrasses
of the Kenyan coast, western Indian Ocean.
The heterocystous phylotypes were closely related to Calothrix and Rivularia,
as well as to unidentified phylotypes associated with seagrasses along the Indian Ocean coast (Uku et al., 2007). Interestingly, one unicellular sequence
from Paper IV grouped with several uncultured cyanobacteria sequences from
tropical oceans, tentatively identified as symbiotic related cyanobacteria earlier
being found as symbionts in sponge (Webb & Maas, 2002).
As seen in Figure 8, the nifH gene phylogenetic analyses also verified that all
sequences analyzed from both Paper III and IV formed potentially new nonheterocystous cyanobacterial clades. These novel non-heterocystous filamentous clades are distantly related to the planktonic Oscillatoriales such as Trichodesmium and Oscillatoria, benthic uncultured phylotypes from Indian
Ocean included Lyngbya majuscula (Lundgren et al., 2003), as well as benthic
phylotype from Pacific Ocean tentatively affiliated to Blennothrix (Díez et al.,
2007). Several other sequences were closely related to the heterocystous phylotypes, formed a separate clade with uncultured phylotypes from seagrasses of
the Indian Ocean and from the Bahamian stromatolites of the Pacific Ocean
(Foster et al., 2009b). The latter clade is in turn closely related to another clade
of heterocystous cyanobacterial phylotypes affiliated to planktonic symbionts
of diatoms (Richelia and Calothrix) occupying waters in the Pacific and Atlantic Oceans (Foster & Zehr, 2006). Indeed, our heterocystous clade may
represent active diazotrophic benthic phylotypes stressing their potential value
as nitrogen-fixers in the benthic marine system here investigated.
Altogether, our morphological and phylogenetic analysis support the suggestion that the majority of the recovered seagrass associated epiphytes are of a
cyanobacterial nature, that a rich morphological diversity is apparent and that a
fraction of these may represents novel unique taxa. Moreover, it is also clear
that a fraction represents potentially diazotrophic cyanobacteria.
Diazotrophs and nitrogenase activity
The light microscopic approach (Paper I, III, and IV) together with the phylogenetic approach in Paper III and IV also provided a detailed account of the
potential diazotrophic community among the cyanobacteria associated to the
seagrass ecosystems investigated here. The diel expression of the nifH gene,
the corresponding biosynthesis of the NifH protein and the in situ nitrogen
fixation activity assays convincingly demonstrated that the seagrass communities contain variable and complex diazotrophic patterns and that this diazotrophic community was dominated by cyanobacteria. In addition, nitrogenfixation (acetylene reduction) rates were appreciable in all seagrasses studied,
36
although they varied depending on season and among the seagrass species
examined, often being higher in Halodule uninervis than in the other genera/species (Paper IV). Hence, these organisms may act as a source of new nitrogen in the seagrass ecosystems of oligotrophic waters.
For instance, in Paper III and IV we showed that the non-heterocystous filamentous cyanobacteria Lyngbya spp. and the heterocystous cyanobacteria Calothrix spp./Rivularia spp. were the dominant diazotrophs in the seagrass phyllosphere investigated. Furthermore, in Paper III, the nifH transcription data
suggested a positive correlation between the diel nitrogenase activity rhythms
and the magnitude of the N fixed and the type of cyanobacteria that dominated.
Maximum nitrogenase activity (Paper IV) occurred when heterocystous cyanobacteria were present. The complex diel nifH gene transcription concurred
with the daily pattern of nitrogenase activity reported in Paper III and IV.
These findings showed that the nifH transcription coincide with nitrogenase
activity, and suggested diazotrophic cyanobacteria as the dominant nitrogenfixers in the seagrass phyllosphere.
The common occurrence and the rates in nitrogenase activity reported in Paper II and III confirmed the importance of seagrass-associated nitrogen fixation
in the coastal areas of the western Indian Ocean. The nitrogen-fixation activity
contributed by the seagrass-associated diazotrophs was predicted to be higher
as compared to activities reported from other ecosystems in the region such as
in relation to mangroves (Lugomela & Bergman 2002; Kyaruzi et al., 2003)
and open intertidal (unvegetated) sediments (Lyimo & Lugomela, 2006). A
summary of the nitrogen fixation activities recorded in Paper II, and a comparison with levels obtained in other marine benthic ecosystems are presented in
Table 1, Paper II (Hamisi et al., 2009).
In addition, a comparison of nitrogenase activities exclusively from the studies of this thesis is presented in Table. 2. As seen from this summary, the
nitrogenase activities were considerable at all sites examined and in relation to
all seagrasses. However, great variations were apparent between seagrasses,
sites, seasons and day/night, with most active fixation being monitored in association with the seagrass at the less human impacted (pristine) site.
37
Table 2: Summary comparison of nitrogenase activities data from this
Thesis: Paper I-IV.
Reference
Sampling site (seagrass genus)
Paper
I
Chwaka, no seaweed farms
(Thalassia)
Chwaka seaweed farms (Thalassia)
Jambiani no seaweed farms
(Thalassia)
Jambiani seaweed farms (Thalassia)
Ocean road (Thalassia)
Mjimwema (Thalassia)
Ocean Road (Cymodocea)
Mjimwema (Cymodocea)
Chapwani, Natural sample (Cymodocea)
After nutrient enrichment, moderate
concentration treatment (Cymodocea)
After nutrient enrichment the highest
nutrient concentration treatment (Cymodocea)
Mjimwema, 2007 (Cymodocea)
Mjimwema, 2008 (Cymodocea)
Paper
II
Paper
IV
Paper
III
Nitrogenase activity
nmol C2H4 g-1 h-1
Time
36 ± 40a
Day
23 ± 23a
13 ± 12a
Day
Day
13 ± 15a
64 ± 40b
124 ± 76b
100 ± 96b
204 ± 76b
46 ± 8c
Day
Day
Day
Day
Day
07.00
70 ± 19c
07.00
8 ± 7d
24.00
358 ± 232
258 ± 139
09.00
21.00
a)
The activity is reported as µmol C2H4 m-2 h-1.
In Paper (II), the original rates were given as nmol nitrogen fixed. The 4:1 conversion factor
was used to convert the activities to nmol of C2H4.
c)
Both the natural population and in the nutrient enrichment experiments with moderate nutrient
concentrations added (moderate in respect to the varied concentrations used within the study),
showed highest activity in the morning (0.700).
d)
The highest nutrient concentration showed highest nitrogenase activity at night time, 2400,
which also indicate that different diazotrophs were involved compared to in the natural samples
and after the moderate nutrient treatments.
b)
Apart from differences in nitrogenase activities between sites as well as between seagrasses, Paper II showed seasonal fluctuations in activity at both the
pristine site (Mjimwema) and the highly impacted site (Ocean Road). A distinct annual pattern was apparent in sediments as well as among the seagrasses.
Higher nitrogenase activity occurred during October to December with the
highest activity in November at both sites and the lowest activity during June
to August (2002–2003) (Fig. 7 in Paper II).
Likewise, the nitrogenase activity in the phyllosphere of the seagrass Cymodocea rotundata assayed in October and November (2007 and 2008) showed a
similar trend in Paper III, with higher nitrogenase activity in November than in
October (Fig. 9). The highest activity encountered was 358 ± 232 and 258 ±
38
139 nmol C2H4 produced g-1 and h-1 in November 2007 and 2008, respectively
(Paper III). Undoubtedly, the nitrogenase activities as well as the cyanobacterial community in general were affected and regulated by the prevailing monsoon winds, affecting for instance currents and temperatures. The south eastern
monsoon (NEM) lasts from December to April, and the north eastern monsoon
(SEM) from June to October, with November and May as intermediate periods.
The highest nitrogenase activities were found in NEM, although this period
was characterized by a low cyanobacterial diversity and percentage cover
(Hamisi et al., 2004). This finding may be explained by the prominent heterocystous cyanobacterial population found in November (Paper III). The lower
cyanobacterial species diversity in November may therefore be compensated
by the potentially higher diazotrophic efficiency of heterocystous cyanobacteria. These findings also show that natural factors may cause shifts in the microbial community, and hence alter the ecosystem function.
Figure 9. Diel nitrogenase activity (acetylene reduction assay) by epiphytes
associated with the phyllosphere of Cymodocea rotundata. The graphs
represent different months (October and November) and years (2007-2008).
The nitrogenase activity is expressed as the ethylene produced per gram
weight of the seagrass per hour; SD (±) is given as bars.
39
Anthropogenic effects
Major disturbances affecting the production of coastal communities are those
linked to physical destruction and/or changes in the water quality, such as via
anthropogenic coastal pollutions, eutrophication and sedimentation. Likewise,
the seagrass associated diazotrophs may potentially be altered by similar external disturbances. The data obtained in Paper I, II and IV clearly showed that
the nitrogenase activity always decreased as a consequence of anthropogenic
activities, such as disturbance related to seaweed cultivation close to seagrass
stands/meadows (Paper I), to sewage discharges (Paper II and IV), and even to
moderate nutrient enrichments (Paper IV).
In paper I, a significantly higher frequency and diversity of cyanobacteria in
seagrass meadows outside the seaweed farms was found. This suggests negative influence on the cyanobacterial composition of such human activities.
Additionally, no heterocystous cyanobacteria was encountered in transects
passing along the seaweed farms at the two sites investigated (Chwaka and
Jambiani, Zanzibar). This decrease may be a consequence of human disturbances via farming related activities. Heterocystous cyanobacteria may be
more susceptible to physical harsh disturbances and to altered nutritional regimes. Moreover, insignificant differences in nitrogenase activity between the
seaweed and non seaweed sites were found at both sites, Paper I.
Furthermore, it was also verified in Paper II that the seagrasses were affected
negatively by increased human pressures. In coastal regions outside the city
Dar es Salaam, a higher biomass was apparent at the less disturbed (less nutrient-rich) site (at Mjimwema; 208 ± 20 g dry wt m–2) compared to the more
eutrophicated site (at Ocean Road; 138 ± 16 g dry wt m–2). These data suggest
that the long term exposures to increased nutrient loads of the latter site affected the seagrass performances more severely. However, data in Paper IV
showed that moderate nutrient administrations did not affect the seagrass biomass and photosynthetic performance during short term exposures, indicating
that seagrasses reactions require higher nutrient loads and longer exposure
times.
Nevertheless, a massive growth of microbial biofilms was apparent in the
mimicked nutrient enrichment experiment of Paper IV. Microscopic observations showed that the cyanobacterial population shifted already under these
milder nutrient addition conditions, with non-heterocystous cyanobacteria being abundant and a few heterocystous species being present. The massive proliferation of non-heterocystous cyanobacteria is in sharp contrast to the cyanobacterial composition on the natural seagrass samples functioning as the starting material. These population changes were also reflected in a significantly
lower nitrogenase activity after higher nutrient enhancements. This illustrates
that the moderate nutrient concentrations used were high enough to inhibit the
nitrogenase activity, while the seagrasses were still not affected.
It therefore is obvious that the potentially more fast growing prokaryotic epiphytes responded considerably faster to disturbances than their more slowgrowing eukaryotic hosts. Hence, changes in microbial populations may be
40
used as ‘early warning’ indicators of ongoing environmental disturbances, of
natural (e.g. climate change) or anthropogenic origin (eutrophication, pollution, etc) in the seagrass ecosystems.
Finally, the findings of the data presented in this thesis no doubt stress the
importance of lessening environmental disturbances in favor of maintaining the
rich natural diazotrophic cyanobacterial biodiversity and their physiological
performances documented here for the more pristine sites, and to practice a
sustainable management of the seagrass ecosystems.
41
Conclusion
In this study we have explored the cyanobacterial diversity and ecological role
as diazotrophs in several tropical seagrass ecosystems. Several specific questions were addressed and the data generated allowed the following more general conclusions to be drawn:
•
The western Indian Ocean seagrass beds support a broad cyanobacterial diversity represented by various morpho- and genotypes.
•
Some of the cyanobacteria are predicted to be novel species, as they
have a specific affiliation to often ‘unknown’ cyanobacteria from similar related under-studied habitats, or geographical locations, only.
•
The diazotrophic cyanobacteria are primarily responsible for the complex diazotrophic patterns of the seagrass ecosystems found, varying in
time and space as well as within the individual seagrass stands.
•
The epiphytic cyanobacteria may constitute a continuous source of
‘new’ nitrogen to the seagrass ecosystem via their nitrogen fixation.
•
Increasing anthropogenic pressures in coastal regions may negatively
affect and shift the cyanobacterial community towards a non-nitrogenfixing community, and thereby alter their proposed ecological function
in the seagrass meadows. Not only differences in cyanobacterial composition and nitrogenase activity, but also difference in seagrass biomass was found as a result of eg. increasing nutrient inflows.
•
Moderate nutrient level increases for shorter periods (comparable to
natural eutrophication pulses) did not affect the seagrass biomass and
their photosynthesis performances, but changes in the cyanobacterial
diversity and nitrogenase activity were observed already at this stage.
•
Changes in microbial populations and functioning may therefore be
used as early indicators of ongoing environmental disturbances due to
increasing anthropogenic activities.
•
The data also stress the importance of lessening anthropogenic based
eutrophication in order to favor and maintain a natural biodiversity of
diazotrophs in coastal ecosystems, particularly the seagrass ecosystems.
42
Way forward
The findings of this work give a deeper insight into the importance of epiphytic cyanobacteria growing within tropical seagrass ecosystems. Still however, many questions remain, such as:
What is the fate of the nitrogen fixed? Is it transferred to the seagrass and if
so, which nitrogen species is transferred, in which quantities and at what rate?
Answers could be achieved by following the nitrogen fixed using the stable 15N
isotope and mass spectrometry. Employing the use of Stable Isotope Probing
(SIP) or Nano Scale-secondary Ion Mass Spectrometry (Nano-SIMS) are other
approaches to determine if the N fixed is incorporated in the seagrass tissue.
What is the nature of the association between cyanobacteria and hosting
seagrasses? Is the interaction specific, and how ‘deep’ is the colonization by
the cyanobacteria? Do any inter- or intracellular penetrations take place? Are
there any seagrass signals released to trigger the colonization? To answer these
questions would e.g. require detailed structural and ultrastructural studies of
some selected model cyanobacterial-seagrass system.
There is also a need of a more coastal/regional specific identification key related to cyanobacteria in the Indian Ocean. Present morphological identifications relay on identification keys of cyanobacteria in remote regions (not always marine) potentially leading to improper identifications.
There is a need to further explore the marine benthic cyanobacterial diversity in relation to other important ecological functions (besides nitrogen fixation)
such as screen for their toxin production and the production of potentially valuable bioactive compounds, if we are to fully understand the contribution of
these fascinating minute organisms.
Additional studies should also integrate more extensive sampling strategies,
isolation (culture dependant) and analysis of whole 16S rRNA gene sequences,
compare with novel marine cyanobacterial genome sequences and metagenomic data, currently becoming available from microbes in the Indian Ocean (via
GOS data).
There is also a need to establish tropical coastal nitrogen budgets to achieve
budgets of the total global marine primary productivity. Additional nitrogen
fixation data, as well as denitrification/anammox data are needed, together
with compilation and extrapolation of available information into local and
global marine models.
In the present study, the presence of toxin producing cyanobacterial strain in
shallow water of the tropic WIO region was not accomplished (due to technical
43
problems). It will therefore be important to perform periodic monitoring activities in this context. This is an important aspect especially in developing countries like those of the African WIO, in which people so heavily depend on natural coastal resources.
44
Acknowledgement
I sincerely wish to express my deepest gratitude and appreciation to my
supervisors Prof. Birgitta Bergman, Dr. Thomas Lyimo and Dr. Beatriz Díez for
their advice, suggestions, comments, constructive criticisms, general guidance
and encouragement led to a successful completion of this study. To me you were
not only the supervisors but co-workers and closest friends ever.
My sincere thanks should go to the SIDA-SAREC for funding this work. I would
also like to acknowledge various agencies (The Swedish Foundation for
International Cooperation in Research and Higher Education, Stockholm Marine
Science Centre (SMF), Western Indian Ocean Marine Sciences Association
(WIOMSA) through MASMA and MARG program, The Swedish Research
Council, K. and A. Wallenberg Foundation and the International Foundation for
Science (IFS)), for providing financial support for analysis and travel grants for
the laboratory visits, courses and scientific occasions during the period of this
study.
I am grateful to the people at the Institute of Marine Sciences (IMS), Zanzibar,
particularly the former Director Prof. Alfonce Dubi, current Director Dr.
Margareth Kyewalyanga, Coordinator of the Sida-SAREC Bilateral project Dr.
Matern Mtolera, for making funds available. I extend my thanks to all staffs at
IMS for logistics supports and field excursion, and students for the company
during my frequent visit at IMS, Asanteni sana!
Many thanks to people at Botan, Stockholm University, especially Dr. Karolina
Ininbergs for being such a good teacher introducing me to the world of molecular
biology, I will not forget the first days at Botan. I wish to extend my thanks to the
previous and present PhD student and visitors in Birgitta’s, Ulla’s, Katharina’s,
Sophia’s, Mats’s group and all of the Botan Department colleagues and staffs for
being so much supportive in both social and academic world, making botan a
colourful place. Tack så mycket!.
I would also like to extend great thanks to my previous and present colleagues
from Tanzania, both Zoology and Botan Department, Stockholm University, for
your nice company, valuable supports and constructive discussion.
45
I would like to extend my sincere appreciation to the administration of The
University of Dodoma (UDOM) for giving me permission to continue with my
study despite of the institution need. I would also like to thank my fellow
employees at the School of Natural Sciences and Mathematics, and UDOM in
general for encouragement especially during the last phase of my study.
I would like to extend my sincerely appreciation to both academic, supportive
staffs and postgraduate student of the Botany Department, Molecular Biology and
Biotechnology Department and the Department of Aquatic Sciences and Fisheries
for the moral support during the whole period of my study. Thanks to all who
helped in logistics and field excursions. Thanks a lot dada’s, kaka’s, shemeji’s,
wifi’s, babu’s, etc for all the memorable moments.
Out of work, I would like to express my special appreciation to the members of
Kilimanjaro Club (Tanzanian in Sweden) for all the gathering, special thanks to
Zam Khalifa, his wife Saadiye and a daughter Leeylat for a warm Tanzanian
kindness. I would also like to extend special thanks to my dearest sister’s,
brother’s, aunties, uncles, friends and neighbours in Tanzania for your moral
support and prayers.
I would like to extend my heartfelt appreciation to my dearest parents, Mama &
Late Baba, for your prayers and love you were always there for me and my
family!
Special thanks and great appreciation to my lovely Husband, whose constant
encouragement, love, care, understanding and unlimited patience supported my
way to success. Thank you very much for being such a good father, taking care of
the family alone for such long! Special thanks to my children, Hannat and
Haroun, and my youngest brother saidi for their patient! Hannat thank you for
counting days, and all the nice stories ‘siri’ on phone, Haroun thanks a lot for not
being tired asking when I will be back!
Last but not least, I would like to convey my thanks to Almighty God, the creator,
to whom I belong, for giving me courage, health and strength!
46
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