Apr 122017

Once we realised that most of our plate of Schistidium ITS2 amplifications had been successful, it was an easy decision to process them all for DNA sequencing. If a higher proportion had failed, we would have had to “cherry pick”, selecting and transferring the successful reactions into new tubes. Every sequencing reaction has a cost, and so deliberately sending a lot of failed reactions through the process, knowing that they won’t generate DNA sequences, is worth avoiding. However, transferring the reactions into new tubes is a meticulous job. In a plate of reactions, there are 96 wells arranged in 12 columns (1-12) and eight rows (A-H).  The first sample is at 1A, then 1B, 1C, 1D, 1E, 1F, 1G, 1H, 2A… all the way to 12H. Imagine if the first failed sample is at 1B and the second at 1G – then we’d transfer 1A to 1A, 1C to 1B, 1D to 1C, 1E to 1D, 1F to 1E, 1H to 1F, 2A to 1G, etc… it’s easy to see that any interruptions can be fatal (this sort of task is why people in the lab sometimes have a “Do Not Disturb” sign stuck on the back of their lab coats). Thus, with only a few failures, we’re better keeping any liquid transfers simple (and manageable using a multichannel pipette).

A 96-well plate on ice

After electrophoresis and visualisation of the Schistidium ITS PCR products under blue, or U.V., light, the next step is to clean up these PCR products. This is where we remove unincorporated primers and dNTPs from the reactions. In our PCRs, we added all the ingredients in excess, so that there was more of everything than the reaction needed. This might seem wasteful, but compared to the costs of the time and plastics that would be required to carefully optimise each reaction, throwing lots in and getting quicker results really isn’t too profligate.

When a PCR is successful, the primer (an oligonucleotide, or short single stranded DNA molecule) binds to the end of the region that is getting copied, and a new strand of DNA is synthesised from the end of the primer, incorporating the primer into the new strand. Thus, primers get used up. The building blocks for DNA synthesis, the nucleotides (As, Ts, Gs and Cs), are the dNPTs (deoxynucleoside triphosphates) that we added to the PCR reaction, again in excess. So even when we’ve built a lot of copies, we still have some primers and dNTPS that weren’t used up.

This didn’t matter when it came to looking at the results on the gel, but does become important in our next reaction, the sequencing PCR. In the sequencing PCR, we add just a single primer, and we add a very precise blend of nucleotides. So we don’t want to carry over primers and dNPTs from the previous reaction. There are several ways of removing these.

The cheapest is through ethanol precipitation of the synthesised DNA, where the unincorporated primers and dNTPs stay in solution and are thrown away. This is less easy to scale up to plates, and the moment where you turn it all upside down and toss the liquid out is rather worrying – the cost of plates of subsequent sequencing failure if the DNA pellets were lost is huge, and the pellets tend not to be visible.

For several years we used a column-based approach, a bit like our DNA extraction method, where the PCR product is bound to a membrane, and the unincorporated primers and dNTPs are flushed off it, before the PCR product is eluted back into solution.

However, the method we now use most regularly involves a combination of two enzymes, Eco1, and Shrimp Alkaline Phosphatase (SAP), which work elegantly in combination, and which involves methodology that is easily scalable for working with plates and multichannel pipettes. The Eco1 enzyme digests single-stranded DNA, so it cuts the primers up into individual nucleotides. The SAP enzyme dephosphorylates the nucleotides (the dNTPs) so that they are no longer functional. Thus after using these enzymes, although nothing has been physically removed from the tubes, the unwanted reagents have been rendered unusable. For convenience, we buy in a commercial combination of the two enzymes called ExoSAP-IT™, and add a small amount of that to our reactions (in this case, around 1 μl ExoSAP-IT to about 15 μl of PCR product).

The enzymes work best at 37ºC, and so the reactions were put on a heating block for 15 minutes. The enzymes are killed at a higher temperature, and so the next step was to heat the reactions to 80ºC for 15 minutes, to make sure that no viable enzymes are left to interfere with the sequencing reaction. After that, the plate of cleaned Schistidium ITS2 amplicons was left in the fridge until it was needed for the sequencing reactions.


Links to reports on Moss diversity in an artificial landscape, an EU Synthesys Access project with Dr Wolfgang Hofbauer at RBGE:

Apr 112017


Gel loading in Lab 33

Once the polymerase chain reaction is over, it’s time to Run The Gel; this is make-or-break time, when we find out if our PCR amplification has actually worked.

The first step is to prepare and pour the gel. The gel is a 1-2% mix of agarose in a salt-containing buffer solution (for us, this is usually 1x TBE); the agarose is dissolved into the buffer using heat, in a microwave, and a bit of stain is added. We use about 5 μl of SYBRsafe for a 100 ml gel. The SYBRsafe binds to DNA and fluoresces under blue light, so we can see if we have produced amplified DNA.

After the agarose gel has cooled down to a comfortable temperature (so we don’t risk warping our ridiculously expensive plastic gel set-up with it!) we pour it onto a plate, and add a “comb” which sits in the gel and around which it will set, forming lines of small oblong holes in the gel’s upper surface into which we will later pipette our PCR product.

Setting up the loading buffer and PCR product in Lab 31

While waiting for the gel to set, we mix a small amount of the Schistidium ITS PCR product (3-5 μl) with a little loading dye (1-2 μl). The loading dye serves two purposes: it contains glycerol, which is rather dense and helps the liquid drop into the holes (wells) in our gel, and it also contains a colour (in this case, bromophenol blue), which means that we can see what we’re doing – dropping a colourless liquid through another colourless liquid into a colourless hole would otherwise be rather more an act of faith!

Using a manual multichannel to load the PCR product/loading solution

When the gel has set hard, we lift off the comb, taking care not to rip the gel (which is rather more brittle than gelatine-based jellies), and put the gel into a gel tank, covered with a thin layer of the TBE buffer that was used to make the gel. Then it’s time to load. Down one side, we put 3.5 μl of a “ladder”, a bought-in product containing DNA fragments of known lengths and quantities, which will act as a standard for the gel and let us estimate the sizes of any PCR products that show up. Adding the ladder, in which we know there is DNA, is also like a “positive control” that we’ve made our gel properly – if nothing at all shows up in the image of the gel, then it’s not proof that our PCR reaction failed, but that there was something wrong with our gel.

With the gel set-up that we are using for our plate of 96 Schistidium amplifications, we can load 8 samples at a time using one of the multichannel pipettes. For this, instead of our usual electric multichannels, we use a manual one so that we have more control over the rate of dispensation, and so we can stop if some of the liquid is not going in the right place: it’s quite a precarious operation!

Running our gel

Because the sugar-phosphate exerior of a DNA molecule has a negative electric charge, DNA molecules migrate through an electric field towards the positive electrode, and so if we apply a current to the gel, we can get the DNA to move through it. The filtering effect of the set agarose in the gel means that the DNA molecules migrate at a rate proportional to their size, with short molecules moving quickly, and longer ones more slowly. Generally, 30-45 minutes at 80 volts is enough to see if our PCR is successful.

After electophoresis, the gel is placed on the transilluminator

After the gel electrophoresis, the gel is carefully lifted out of the buffer, off the gel tank, and placed on a plate in a light box. (At this point with larger thinner gels like the one we ran for the Schistidium PCR products, it’s quite easy to break the gel.) Because the shape of the gel and of the light box are different, these gels need to be cut in two. Luckily, the halves fit nicely side by side in a single image.

Our gel beneath the filter

An orange filter is added, and when it’s turned on, the transilluminator shines with blue light that will make any DNA stained with SYBRsafe fluorescent, at which point a digital image is taken of the gel.


The first image of our Schistidium PCR products

To see what’s on the gel, we use an automated setting, where the computer choses the best exposure for the image. However, a bit of subsequent cropping and fiddling with the image itself can make for a far better gel picture. And the really good news for us is that, looking at our gel, it is clear that most of our Schistidium DNA extractions were successfully amplified for the ITS2 region in our PCR reaction!

The wells can be seen at the top of each row, as darker shadows; for the purposes of this image, the DNA has migrated towards the bottom of the screen. For only about seven of the 96 reactions is there no visible band of PCR product on the gel, and in the vast majority of cases a clear bright single band is present. (The fuzzier band that is sometimes visible a little further down than the ITS product, especially for the second last sample on this gel, is primer-dimer, an artifact caused during PCR, and not the amplified product we want; it’s far far shorter than the region we’re working with, as can be seen by comparing it with the size of our 1 kilobase DNA ladder).

Cropped and adjusted gel image picture, with a ladder on the right hand side of every third row

Links to reports on Moss diversity in an artificial landscape, an EU Synthesys Access project with Dr Wolfgang Hofbauer at RBGE:

Apr 062017

PCR set-up with defrosting reagents in Lab 32

After we extracted a plate’s worth (12 columns by 8 rows, or 96 samples) of Schistidium DNA, the next step in our process is to copy a preselected part of that DNA, using the Polymerase Chain Reaction (PCR). For this study with Wolfgang, we are copying a region of nuclear DNA known as the Internal Transcribed Spacer 2 (ITS2). We set up the reactions in a laminar flow hood, which blows clean air down onto the work surface, keeping everything as clean as possible. Most of the reagents are kept in a -20ºC freezer between uses, so these get set out to defrost before use (time for a coffee break!). The DNA polymerase enzyme, taq, on the other hand, is stored in glycerol so doesn’t freeze at -20ºC; while we’re using it, we keep it on ice so it stays cold.

The reaction components are added into a single “Master-Mix” tube, which includes water, buffer, Magnesium, enhancing additives, short oligonucleotides (primers), and the taq enzyme. Small (19 μl) aliquots of this master mix are then added to a new 96-well plate, and 1 μl of DNA from the extraction plate is transferred across into each of the 96 reactions.

Using the multichannel pipette to transfer DNA

Once the DNA has been added, the plate is sealed (this time with a clear plastic film that sticks firmly in place when heated), briefly spun in a centrifuge to make sure all the reagents are mixed together at the bottom of the 96 plastic wells, and transferred into one of our Thermocyclers, or PCR machines.

PCR plate in Lab 32 thermocycler

The plate sits in a metal block which rapidly “cycles” up and down in temperature, following a predetermined programme. For the gene region that we are copying here, the programme first heats the block to 95ºC for 4 minutes, then starts a cycle of 94ºC for 1 minute, 55ºC for 1 minute, and 72ºC for 45 seconds, repeated 30 times. When the block is at 94ºC, the double-stranded DNA is pulled apart into single strands; when it’s cooled to 55ºC the primers stick on and the taq initiates making copies of the ITS2 region, and when it is heated up again to 72ºC, the copies get completed, before they’re pulled apart again when the block heats up to 94ºC and the cycle starts again…

The thermocycler screen showing the ITS2 PCR programme

A couple of hours later, the reaction is complete, and at this point we HOPE that all the 96 wells in our plate contain millions of copies of the ITS2 DNA molecule from the Schistidium extraction that was added in each case. However, in order to see if any of it has actually worked, we need to stain and visualise the DNA, and for this, we have to run a gel, a process that will be the subject of the next installment.



Links to reports on Moss diversity in an artificial landscape, an EU Synthesys Access project with Dr Wolfgang Hofbauer at RBGE:

Dec 022016
Some of the herbarium collections of Marchantia held in the RBGE herbarium

Some of the herbarium collections of Marchantia held in the RBGE herbarium

Many new species are already included in natural history collections around the world, it’s just that nobody has yet got around to examining the material, recognising that it represents something novel, and publishing a name for it. Sometimes these new species are filed under the epithet of a similar named species, sometimes they’re just filed under the genus name with other collections that have not been identified to species, and sometimes they have been annotated to recognise that they’re probably distinct from all the species that have already been described, e.g., as “sp. nov.

David Long has made a huge number of plant collections from around the world in his 40-plus year botanical career, with many of these collections not yet fully examined. Some of this material is being mined for DNA sequencing projects at RBGE, and for some of our key plant groups, as well as sequencing well-identified material, we are also sequencing plants that have not been assigned to species. Molecular lab work is fast compared to close morphological studies of multiple plant specimens; this can therefore speed up the processes of traditional taxonomy, by allowing it to focus on things that are obviously distinct.

One lineage that David Long is particularly involved with, and that remains one of our key plant groups, is the complex thalloid liverworts. Some of our sequencing work has involved Marchantia, which made Xiang et al.‘s recent description of a new species in the genus, Marchantia longii, particularly interesting. In the last few days, the DNA sequences that were included in the paper were made publicly available on the NCBI site, GenBank. One of the regions that was sequenced by Xiang et al., the plastid-encoded RuBisCo Large subunit gene rbcL, was also included in our study, and so I was able to put the two data sets together, and see how the new species fits into our phylogenies.

The results are interesting: When Xiang et al. named M. longii, they did so in part because the area that the plant came from, in northwestern Yunnan, is one in which David has been very active. In fact, at RBGE we had already generated DNA sequence data from nine accessions of Marchantia that David had collected there. I was delighted to find that two of these accessions (collections Long 36155 and Long 34642), which had been filed in our collections without a specific epithet, are an exact genetic match to Marchantia longii. It seems that David really does have an affinity for the plant, having gone out and found some even before it was named for him!


Long’s Marchantia

A rapid phylogeny of Marchantia, from the RBGE collections. II. Illuminating our sampling

A rapid phylogeny of Marchantia, from the RBGE collections. I. Sampling

Sep 122016
Photo 11-09-2016, 12 14 22

The tiny liverwort Colura calyptrifolia (photographed with an iPhone and a x20 handlens!)

Colura calyptrifolia (or to give it its appropriately creepy-sounding common name, the Fingered Cowlwort), is one of our most fascinating UK liverworts. Absolutely tiny (the leaves are about a millimetre long and whole plants often only 2-3 mm), it is heavily modified from the basic leafy-liverwort body plan, the leaves formed into inflated sacs like miniscule balloons with pointed “beaks” at one end. These tiny sacs have even tinier trapdoor-like flaps that only open inwards, allowing them to capture ciliate protozoa and other microscopic creatures (conclusively observed in another species of the same genus). It’s not yet certain if the liverwort gains nutrients from “swallowing” these animals, although this might be a reasonable hypothesis given the similarity of the mechanism to that found in much larger carnivorous plants such as bladderworts.

Photo 11-09-2016, 12 14 20

The leaves of Colura are modified into tiny ballon-like sacs that trap small animals

This colony was spotted yesterday in Anglezarke, Lancashire following the British Bryological Society (BBS) AGM in Manchester. Populations of Colura have undergone a spectacular expansion over the last 10 or 20 years, particularly in conifer plantations where they occur as tiny epiphytes. Previously the plant was rather rare, occurring mostly on rock in humid gullies and restricted to the wetter areas of the far west. This sighting was on the trunk of a willow at the edge of a reservoir.

Something else that has changed rapidly over the last 10 or 20 years is the ease with which small things can be photographed and shared. These pictures were taken simply by pointing the camera of an iPhone through a x20 handlens (the latter much cheaper than an iPhone and even more useful!),  and if we had wished could have been made available online instantaneously. As poorly-known biodiversity is increasingly threatened globally, should we be making better use of cheap imaging and real-time networking of expertise to facilitate species discovery and monitoring?



Barthlott, W., Fischer, E., Frahm, J.-P. & Seine, R. (2000). First Experimental Evidence for Zoophagy in the Hepatic Colura. Plant Biology 2(1):93-97. 

Blockeel, T L, Bosanquet, S D S, Hill, M O and Preston, C D (2014). Atlas of British & Irish Bryophytes. Pisces Publications, Newbury.

Sep 092016
Sphaeropcaros texanus photographed by David Long (Long 33162)

European material of Sphaerocarpos texanus, photographed by David Long (Long 33162)

The Sphaerocarpales (or “Bottle Liverworts”) form a very distinct group in the complex thalloid liverworts, with ca. 30 species in five genera: originally the group just included Geothallus (monospecific), Sphaerocarpos (8-9 species) and Riella (ca. 20 species), with two more monospecific genera, Austroriella and Monocarpus, added within the last few years. All five genera have very unusual, and highly reduced, thallus morphologies. With the exception of Monocarpus, they also all enclose their sex organs (or gametangia – the antheridia and archegonia) in inflated flask-shaped bottles (as can be seen in the accompanying photograph). This feature sets them apart from all other liverworts. All of them are adapted to extreme habitats, including arable fields, hot arid regions, seasonal lakes and pools, and salt pans.

A worldwide revision of the second largest genus of the group, Sphaerocarpos, is over 100 years old (Haynes 1910); other revisional work focuses on individual geographic areas, including South Africa (Proskauer 1955), North America (Haynes & Howe 1923, Frye & Clark 1937, Schuster 1992, Timme 2003), California (Howe 1899), Europe (Reimers 1936, Müller 1954), and France (Douin 1907). No revisions have been made for large areas including Australia, Asia and South America, and most of the work predates any DNA-based concepts of plant identification or species relationships. Bringing the taxonomy of Sphaerocarpos into the 21st century, Dr Daniela Schill spent 18 months (2007-2009) at RBGE on a Sibbald Trust-funded project to compile a world-wide taxonomic revision of the genus. Two field expeditions fed into the project, with Dr David Long collecting European species in Portugal in April 2007, and Daniela collecting North American species in California in March 2008 (funded by the Peter Davis Expedition Fund).

Spore SEMs of Sphaerocarpus drewiae, taken by Daniela Schill

Spore tetrads of Sphaerocarpos drewiae, SEMs taken by, and plate prepared by, Daniela Schill

Daniela’s work is based on morphological and anatomical characters, including spore characters that she observed using Scanning Electron Microscopy (SEM). Her aim has been to produce identification keys to the species, species descriptions, species lists, synonyms, botanical drawings, distribution maps, and ecological, nomenclatural and taxonomical notes. Although the study is not yet published, much of it, including SEM plates for spores from the ca. 9 different species (as seen on the right), is complete.

In parallel, RBGE staff have also been sequencing multiple accessions of all available Sphaerocarpos species, producing data that has helped inform some of Daniela’s taxonomic decisions, and that also allow us to generate a stand-alone phylogeny for the genus.

This research will lead to some taxonomic changes. For example, European Sphaerocarpos texanus plants differ from American S. texanus, both in their DNA sequences and in their spore characters, and so they are likely to be considered a separate species. Furthermore, European Sphaerocarpos michelii material includes three different forms based on spore characters; these are also confirmed by molecular research, and may be recognised at or below the rank of species.



Cargill, D.C. & J. Milne. 2013. A new terrestrial genus and species within the aquatic liverwort family Riellaceae (Sphaerocarpales) from Australia. Polish Botanical Journal 58(1): 71-80.

Douin R. 1907. Les Sphaerocarpus français. Revue Bryologique 34(6): 105-112.

Frye T.C. & L. Clark. 1937. Hepaticae of North America. University of Washington Publications in Biology 6: 105-113.

Haynes C.C. 1910. Sphaerocarpos hians sp. nov., with a revision of the genus and illustrations of the species. Bulletin of the Torrey Botanical Club 37(5): 215-230.

Haynes C.C. & M.A. Howe. 1923. Sphaerocarpales. North American Flora 14: 1-8.

Howe  M.A. 1899. The hepaticae and anthocerotes of California. Memoirs of the Torrey Botanical Club 7: 64-70.

Müller K. 1954. Die Lebermoose Europas. In: Rabenhorst’s Kryptogamenflora von Deutschland, Österreich und der Schweiz. 3. Auflage. Volume VI. Part 1. Leipzig, Akademische Verlagsgesellschaft Geest & Portig K.-G., Johnson Reprint Corporation (1971), New York, London.

Proskauer J. 1955. The Sphaerocarpales of South Africa. The Journal of South African Botany 21: 63-75.

Reimers H. 1936. Revision des europäischen Sphaerocarpus-Materials im Berliner Herbar. Hedwigia 76: 153-164.

Schill D.B., L. Miserere & D.G.Long. 2009. Typification of Sphaerocarpos michelii Bellardi, S. terrestris Sm. and Targionia sphaerocarpos Dicks. (Marchantiophyta, Sphaerocarpaceae). Taxon 58(2): 638-640.

Schuster R.M. 1992. Sphaerocarpales. In: The hepaticae and anthocerotae of North America V. Field Museum of Natural History, Chicago: 799-827.

Timme S.L. 2003. Sphaerocarpaceae. In: Bryophyte Flora of North America, Provisional Publication.


Sep 062016

University of Edinburgh/RBGE student David Bell, studying for the Masters degree in the Biodiversity and Taxonomy of Plants; thesis submitted August 2009.

Supervisors: Dr David Long and Dr Michelle Hart.


David used plastid DNA barcode markers rbcL (from 34 accessions) and psbA-trnH (from 36 accessions) to look at the four species of Herbertus in Europe, H. aduncus subsp hutchinsiae (British Isles, Norway and Faroes), H. stramineus (British Isles, Norway and Faroes), H. borealis (Scotland and Norway) and H. sendtneri (European Alps).

In addition to the four recognised taxa, David’s study identified a fifth species, later named as H. norenus, that occurs in Norway and the Shetland Isles.

A paper based on David’s MSc thesis work was published in Molecular Ecology Resources in 2012.

Herbertus norenus, photographed by David Long

Mixed sward including Herbertus norenus, photographed in Shetland by David Long


Bell et al. 2012, MER



Other student projects at the Gardens:

Student projects at RBGE: DNA barcoding British liverworts: Lophocolea

Student projects at RBGE: Barcoding British Liverworts: Plagiochila (Dumort.) Dumort.

Student projects at RBGE: Barcoding British Liverworts: Metzgeria

Aug 122016

University of Edinburgh Biotechnology student Kenneth McKinlay’s 4th year honours project, 2013. Supervisors: Dr David Long, Dr Laura Forrest

David Long and Kenneth checking out the Lophocolea on a decaying log in the Scottish Borders

David Long and Kenneth check out Lophocolea on a decaying log in the Scottish Borders

Kenneth barcoded all six species of British Lophocolea, L. bidentata, L. bispinosa, L. brookwoodiana, L. fragrans, L. heterophylla and L. semiteres, attempting to get data from three plastid regions (rbcL, matK, psbA-trnH) and one nuclear region (ITS2). The data generated from the rbcL and psbA-trnH regions was effective in discriminating between all the species sampled; however useful data were not obtained from matK or ITS2.

Genetic markers:

1. rbcL: bidirectional sequence data was generated for 38 accessions.

2. matK: amplification was not successful with the primer sets used (LivF1A, LivR1A).

3. psbA-trnH: bidirectional sequence data was generated for 40 accessions.

4. ITS2: although PCR amplification was successful for 35 accessions, the low quality of many of the sequences generated, and the presence of clear heterozygous positions in sequence data from some accessions, made this data set problematic to analyse, so it was excluded from the study.

Lophocolea bispinosa vice county 98 Long 4725

Lophocolea bispinosa vice county 98, Long 4725; photographed by David Long

Lophocolea semiteres vice county 98, Long 0578

Lophocolea semiteres vice county 98, Long 0578; photographed by David Long









Species and trees:

Distance tree generated using rbcL barcode sequence data for UK Lophocolea accessions

Distance tree generated using rbcL barcode sequence data for Lophocolea accessions, rooted on Chiloscyphus

L. fragrans – all accessions were genetically uniform, forming a monophyletic group.

L. heterophylla – although there was a little genetic variation, again, accessions of this species formed a distinct clade for both rbcL and psbA-trnH.

L. semiteres & L. brookwoodiana – these formed a single clade. All the accessions of L. semiteres (including material from the UK and Belgium) were genetically uniform, while two different genotypes were observed for L. brookwoodiana. While L. semiteres is known to be an introduced species in the UK, it’s possible that the three different genotypes in this clade represent separate introductions.

L. bispinosa – species formed a single genetically uniform group; this nests within a L. bidentata grade.

L. bidentata – accessions of this widespread and common species formed a grade, with three genetically distinct groups. One of these groups may represent L. cuspidata, a species that was sunk into L. bidentata by Bates and Walby in 1991, due to a lack of consistently distinguishing morphological characters. The results of this study suggest that a recircumscription of L. bidentata, “probably the commonest leafy liverwort in the British Isles” (Hodgetts, 2010), is required.


Related Posts

Student projects at RBGE: DNA barcoding British liverworts: Lophocolea

Student projects at RBGE: Barcoding British Liverworts: Plagiochila (Dumort.) Dumort.

Student projects at RBGE: Barcoding British Liverworts: Metzgeria

Student projects at RBGE: DNA barcoding of the leafy liverwort genus Herbertus Gray in Europe and a review of the taxonomic status of Herbertus borealis Crundw.



Jul 192016
Decaying wooden fence, between concrete poles, Kufstein, Austria

Decaying wooden fence, between concrete poles, Kufstein, Austria

Recently in Kufstein, the home of Austrian bryologist Wolfgang Hofbauer, the demolition of an attractive old building and clearing of trees and other plants from the land, leaving a bare gravel patch used as a parking space, did have one interesting outcome: The new clearing led Wolfgang’s eye to a decaying wooden fence between concrete posts. Both the posts and the fence are partly covered in bryophytes, but among them, Wolfgang was very surprised to find the moss Schistidium growing on the old wood as well as on the concrete.

Schistidium on fence post, Austria

Schistidium on fence post, AustriaIn the bryological literature, the only reference to the plant growing on wood is a rare occurrence of Schistidium apocarpum, on lime-impregnated tree bark. The situation in this Kufstein parking lot seems unique, with at least two different species of Schistidium on the wood (although species identification is ongoing). Other more typical residents of old wood, which are also present, include Leucodon sciuroides, Orthotrichum affine and Hypnum cupressiforme. However, the unique assemblage is unlikely to last, as the climatic regime at the place will have changed following the removal of the trees, and the newly exposed rotten fence will probably soon be replaced.

Schistidium on fence post, Austria

Schistidium on fence post, Austria

Meanwhile, however, we wonder if similar unlikely assemblages of mosses are being observed elsewhere, and if there is an explanation for any potential changes in habitat?



Botanics Story and images provided by Wolfgang Hofbauer


Related literature

Wolfgang Karl Hofbauer, Laura Lowe Forrest, Peter M. Hollingsworth, Michelle L. Hart. 2016. Preliminary insights from DNA barcoding into the diversity of mosses colonising modern building surfaces. Bryophyte Diversity and Evolution 38(1).

Sam Bosanquet. 2010. Schistidium species reports, in: Atherton, Bosanquet & Lawley, Mosses and Liverworts of Britain and Ireland a field guide, British Bryological Society.

In plain sight – the mosses that grow on British walls. http://stories.rbge.org.uk/archives/19957

Hidden diversity in unexpected places – moss growth on modern building surfaces. http://stories.rbge.org.uk/archives/17489


May 132016

Plant diversity does not have to be far-flung and exotic to be worth studying; even within Scotland, there are unanswered questions about plant distributions. Growing in our towns and cities, sharing our walls and pavements, there are bryophytes, tiny mosses and liverworts. We pass these every day, step over them, walk past them, hardly noticing that they are there. Miniature ecosystems form in the mosses that grow in the mortar between our bricks, or cling to cement surfaces of our bridges, and yet, partly because they are so commonplace, we don’t usually see them at all. And we have amazingly little understanding of exactly which species are involved, or where they have come from.

Recently, we looked at plants of the common moss Schistidium to find out exactly which species grow on artificial surfaces, like cement, walls and roofs (Hofbauer et al. 2016). Our study included plants from different geographic areas, with many plants collected in Germany and Austria, where Wolfgang Hofbauer, the lead researcher on the study, works and lives. However, a small subset of the plants were collected in the UK, and so also form part of the Royal Botanic Garden Edinburgh’s “Barcoding the British Bryophytes” project. Of 29 Schistidium plants collected in the UK, nine were collected on natural surfaces, like boulders and cliffs, and 17 were collected on artificial surfaces, like walls and roofs (for three accessions we don’t have a record of what kind of surface they were growing on).

Schistidium, photographed by Wolfgang Hofbauer

Schistidium, photographed by Wolfgang Hofbauer

These UK moss samples probably belong to eight species, Schistidium crassipilum, Schistidium pruinosum, Schistidium elegantulum, Schistidium strictum, Schistidium papillosum, Schistidium apocarpum, Schistidium trichodon and Schistidium dupretii, with three of the species, Schistidium crassipilum, Schistidium elegantulum and Schistidium apocarpum, having been collected from both natural and man-made surfaces.

A diagram of genetic relationships between the plants we sampled is shown below.

Schistidium crassipilum – we found three distinct genetic types within this species, which may belong to different species or subspecies. Schistidium crassipilum is known to be common on man-made habitats across Britain and Ireland, and we have collected it on bricks, cement, and even roofs as well as on natural substrates.

Schistidium pruinosum – only one of the moss plants in the study, collected in the Pentlands near Edinburgh, belonged to this species. It’s not known from many collections in the UK, although this may just be because the plants are often overlooked or misidentified, rather than that they are rare.

Schistidium elegantulum – this has been reported from natural and man-made habitats to the south and west of Britain. However, in our study, we have found it growing in the east, on cement in East Lothian and Midlothian, as well as in some more traditionally westerly locations in Scotland.

Schistidium papillosum – only one of the moss plants in this study, collected from limestone in Craig Leek, near Braemar, probably belongs to this species.

Schistidium strictum – again, only one of the plants in our study, collected in Dumfries on rocks, probably belongs to this species.

Schistidium papillosum is sometimes considered to be the same species as S. strictum (e.g. by AJE Smith 1978, The Moss Flora of Britain and Ireland, Cambridge University Press, but not by Bosanquet 2010, p. 515, in Atherton, Bosanquet & Lawley, Mosses and Liverworts of Britain and Ireland a field guide, British Bryological Society), although we did find genetic differences between the two plants that we sampled, consistent with their recognition as two separate species.

Schistidium apocarpum – this is one of the more common Schistidium species, and known to occur on natural and man-made surfaces; we sampled several plants from this species, growing on walls and rocks.

Schistidium trichodon – described as “a rare upland calcicole” by Sam Bosanquet (2010, p. 515, in Atherton, Bosanquet & Lawley, Mosses and Liverworts of Britain and Ireland a field guide, British Bryological Society), both our collections matched the reported habitat, growing on limestone, in Clova and Feith, Scotland.

Schistidium dupretii – we only sampled a single British accession of this species, another rare calcicole, which had been collected at Ben Lawers.

We are still far from having full records of how much genetic diversity there is in Schistidium in the British Isles. Partly because our previous work has focused on mosses on man-made surfaces, we don’t yet have any data for several other species that have been reported from Britain and Ireland (Bosanquet 2010, in Atherton, Bosanquet & Lawley, Mosses and Liverworts of Britain and Ireland a field guide, British Bryological Society). These include Schistidium maritimum (reportedly usually northern and western, in coastal locations), Schistidium rivulare (commonly around water, particularly fast-flowing rivers), Schistidium platyphyllum (another species that grows near rivers), Schistidium agassizii (rare, aquatic and probably often overlooked), Schistidium flexipile (very infrequent, with only one record from recent years), Schistidium robustum (an uncommon upland calcicole), Schistidium confertum (an uncommon upland species), Schistidium frigidum (yet another uncommon reportedly upland species) and Schistidium atrofuscum (a rare moss, only recorded for the UK in the central Highland area).

But at least we are now starting to get a better picture of the mosses that share our towns and cities!


UK Schistidium accessions, parsimony analysis of ITS data with bootstrap support above branches

UK Schistidium accessions: parsimony analysis of nuclear ITS DNA sequence data, with bootstrap support above branches


This work was supported by EU SYNTHESYS project (http://www.synthesys.info) gb-taf-3881.
Thanks are also due to David Long for providing many of the specimens.

Wolfgang Karl Hofbauer, Laura Lowe Forrest, Peter M. Hollingsworth, Michelle L. Hart. 2016. Preliminary insights from DNA barcoding into the diversity of mosses colonising modern building surfaces. Bryophyte Diversity and Evolution 38(1)

Sam Bosanquet. 2010. Schistidium species reports, in: Atherton, Bosanquet & Lawley, Mosses and Liverworts of Britain and Ireland a field guide, British Bryological Society.