Summer in Antarctica is characterized by continuous high
solar irradiance, water column stratification and intense primary production
from phytoplankton. As winter moves in, sea ice forms and the marine habitat is
plunged into darkness; the lack of sun light and lowering temperatures cause
the water column to mix and primary production from photosynthesis virtually
ceases. Previous studies have identified that these seasonal changes in
conditions correspond to a drastic change in microbial abundance and activity,
as well as a shift in microbial diversity. To date research efforts to identify
the main players in the different microbial communities have been limited. Furthermore
there has been no specific objective to analyse the metabolic capabilities of
the different communities. In particular there is a lack of information on
winter communities due to the harsh conditions and thick sea ice which hampers
sampling efforts. The objective of the paper reviewed was therefore, to better
define the community diversity and genome-encoded capabilities of Antarctic
bacterioplankton in both winter and summer climates.
Grzymski et al., (2012) use a bombardment
of metagenomic techniques in order to gather the desired information about each
microbial community; allowing them to observe data on organisms which may not be
culturable. Genetic parameters directly measured included subunit ribosomal RNA
(SS rRNA) sequences as well as genomic end sequences; this was achieved using a
standardised DNA/RNA extraction kit and following instructions from the Joint
Genome Institute. The genomic information was then subjected to a variety of
phylogenetic statistical analyses and comparisons using genomic databases, allowing
data such as predicted genome size, GC content and specific functional genes to
be investigated.
The paper identifies many different clades in each community
using the SSrRNA which enabled them to produce phylogenetic trees. Overall it
was found that the winter community had a significantly higher phylogenetic and
functional diversity. The general unique features of the winter community were
the presence of chemolithoautotrophic bacteria and archaea (who obtain energy from the oxidation of inorganic
compounds and carbon from the fixation of carbon dioxide). Interestingly
the summer community showed no archaea at all. The authors were able to come to
these conclusions from a wide range of interesting and detailed data, not all
of which can be summarised here. I found the specific functional
gene search particularly interesting; this showed that there is a greater
metabolic and functional diversity in winter. For example, the winter community
contained more post-translational modification genes, indicating that a higher amount
of protein re-folding occurs in winter. This is likely due to protein damage in
the more stressful lower temperatures and oligotrophic condition.
The observation of high levels of chemoloithoautotrophy in
winter is novel in Antarctic waters. Given the huge size of the Southern Ocean
this discovery, and its potential inferences, is particularly important. More
work is needed in order to find out if this process is widespread in the
Southern Ocean as it may be an important carbon sink which has previously been unaccounted
for in carbon budget studies. It may also go some way to explaining anomalies in
inorganic nitrogen content, possibly changing the way we understand the global
carbon and nitrogen cycles.
I decided to review this paper in order to better understand
metagenomic techniques and their applications following my earlier confession
that I didn’t have a good grasp of it all. I feel this paper communicated their
huge array of results clearly which helps overall understanding and I strongly
recommend any other bloggers in the same position give this a read.
Grzymski, J. J., Riesenfeld, C. S.,
Williams, T. J., Dussaq, A. M., Ducklow, H., Erickson, M., Cavicchioli, R., et
al. (2012). A metagenomic assessment of winter and
summer bacterioplankton from Antarctica Peninsula coastal surface waters. The
ISME journal, 6(10), 1901–15.
Hi Vicky, you said they found no Archaea at all. Is it possible that their methods of collecting samples were not sufficient? For instance, I would describe a good sampling tecnique as collecting from a variety of different depths to achieve an overall image of the composition. This is especially important when considering photosynthetic organisms, for example, such as in this paper, that require sunlight and would therefore flourish near the surface.
ReplyDeleteHi Megan, thanks for your comment. I'm glad you've brought this point up as it was something I alluded to in the blog and was going to explore more but I ran out of words. Firstly, this study focussed their investigation on surface waters and were therefore unlikely to prioritises depth profiling, perhaps I should have made this clearer when stating their aims. Secondly, in winter the harsh conditions control what sampling can occur, which isn't ideal, but the authors do recognise this. The winter samples were taken directly from the stations sea water intake which is 6m deep and 16m from shore, whereas the summer samples were taken 500m offshore at a depth of 10m using a submersible pump.
ReplyDeleteAs for the lack of archaea, the authors found no evidence of PCR amplified DNA and therefore concluded that they were not present. Given that this finding was for the summer community where sampling difficulty isn’t an issue they concluded this to be correct as it also concurs with previous reports (Murray et al. 1998).
Whilst the authors do recognise that sampling is limited to what the climate permits, I do think that the paper would be improved by using a better sampling regime. Also, as I mentioned in the blog, more work is required in order to get a better understanding of the microbial communities in the Southern Ocean as a whole.
Murray AE, Preston CM, Massana R, Taylor LT, Blakis A, Wu K et al. (1998). Seasonal and spatial variability of bacterial and archaeal assemblages in the coastal waters near Anvers Island, Antarctica. Appl Environ Microbiol 64: 2585–2595