Nov 162016
 
Telaranea tetradactyla, photographed by David Long (Long 37778)

Telaranea tetradactyla at Benmore, photographed by David Long (Long 37778)

Murphy’s threadwort (Telaranea murphyae) has had a singular position in the British flora. The species was described by renowned bryologist Jean Paton in 1965, from plants collected in the south of England. It’s a tiny leafy liverwort that is found in only four locations, at Tresco and St Mary’s on the Isles of Scilly, Branksome Chine, Poole in Dorset and Alum Chine, Bournemouth. Murphy’s threadwort has always been known to be an alien species in our flora, and yet because it’s never been found elsewhere, the sole responsibility for conserving the species lay with the UK. Being non-native, however, it was not considered a priority for UK Biodiversity Action Plans.

Telaranea tetradactyla from the RBGE fern house, photographed by Lynsey Wilson

Telaranea tetradactyla from the RBGE fern house, growing with Conocephalum conicum; photographed by Lynsey Wilson

Using DNA sequence data from the plant, and comparing it to sequences from other related species, we showed that genetically, the English plants are the same species as a New Zealand plant, Long’s threadwort (Telaranea tetradactyla, synonmy Telaranea longii). Long’s threadwort was already known from several locations in the UK, including inside the fernhouse at RBG Edinburgh, and near the fernery in Benmore. These habitats are not entirely coincidental – the Victorian craze for ferns saw many gardens import living tree ferns from countries such as Australia and New Zealand, with many smaller plants hitching a ride along on their trunks. Today, conscious of plant health issues and the potential transport of pathogens, new plant living collections have to spend time in quarantine before being planted out; past gardeners were less careful, and some of these hitchhikers have subsequently escaped into the local landscape.

Telaranea tetradactyla from the RBGE fern house, photographed by Lynsey Wilson

Telaranea tetradactyla from the RBGE fern house, photographed by Lynsey Wilson

Sinking our UK Murphy’s threadwort plants into the New Zealand species means that any conservation requirements now rest instead with New Zealand, although we can continue to enjoy seeing this diminutive mat-forming liverwort in its select few UK locations.

 

 

Key reference: Porley, R.D., 2013, England’s Rare Mosses and Liverworts. Princeton University Press.

 

 

Villarreal et al. 2014, Journal of Bryology 36(3): 191-199

Villarreal et al. 2014, Journal of Bryology 36(3): 191-199

 

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.

 

References:

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.

 

Aug 302016
 
EDNA label printer

The EDNA label printer in the office

Over the years, many different people have used the molecular laboratories at RBGE, to work on a multitude of projects on a multitude of plants and fungi. Some are staff members who stay for decades, others students who are only in the lab for a matter of months. Every time DNA is extracted and used in a molecular project, the amplified gene regions are processed and then the plastic tubes that they were in are sent for recycling – but the extracted DNA is kept in a DNA bank, in case it is needed for further research. Logistically, managing this DNA can be problematic. Scientists like to use their own numbering systems when they’re working (mine used to be one of the commonest – my initials followed by consecutive numbers, a system which worked perfectly until some of my extractions ended up in the same freezer as extractions by Dr Linda Fuselier), something quick and easy to scrawl onto the plastic tubes. This can link to collection information written in a lab-book, including who collected the plant, what date it was collected, and what country it came from. However, as people move on, and as the years pass, it becomes increasingly difficult to find any particular sample or set of samples, particularly when several sets of people share the same initials – and this is compounded by having to rummage through boxes of frozen DNA samples being kept at either -20° or -80°C. Few places at the Botanics are less pleasant than the dank room that contains our -80°C freezers!

 

Printed labels and EDNA tubes, Lab 32

Printed labels getting stuck onto EDNA tubes, Lab 32

The frustrations associated with rooting through inconsistently labelled DNA collections led Dr Michelle Hart and Alex Clarke, in 2006, to instigate a standardised format for DNA labelling, with samples of DNA identified as part of the RBGE DNA bank and assigned EDNA numbers, the format of which consists of the year the DNA was banked, followed by a multi-digit identification number. For example, the last EDNA number that we have issued is EDNA16-0045851, for DNA extracted from the moss Weissia controversia. Due to uncertainties about institutional databases, in its early years the DNA bank was curated through Excel spreadsheets; this was revamped and upgraded in 2011 to the database that we still use today. Information about the methods and date of DNA extraction, the material’s collector, and the place of collection are all stored and easily retrieved, critical information if the DNA is going to be used to provide data for future publications. The EDNA number stays on all downstream files that are created from the DNA – lab books, raw sequence files, and it is also included as the isolate number in GenBank submissions – meaning that all molecular data generated at RBGE is still valuable after people have moved on and lab books have been mislaid.

 

EDNA tube

A labelled EDNA tube ready for the DNA sample, Lab 32

As to what happens to the actual DNA extraction, long-term storage involves transferring the liquid into a small barcoded and labelled tube in a lockable and numbered 96-tube rack, which will be kept on a labelled shelf in a -80°C freezer. The system is not perfect, however – banking or recovering the DNA samples still involves a trip to our mildewy bank room…

 

Pipelling DNA samples into EDNA tubes, Lab 31

Pipelling DNA samples into labelled EDNA tubes, Lab 31

Aug 232016
 

When people extract DNA in the RBGE molecular lab, we insist that it’s given something we call an EDNA (Edinburgh DNA) number. This links to a database that is internal to RBGE.

evilednaThe EDNA number is used for all internal molecular lab processes – it’s written on the tube of DNA, used to refer to the sample in lab books, and part of the file name for all DNA sequences that are generated from that sample. Using this standard system across all projects means that we can keep track of what DNA we have, we can store it in a way that makes it relatively easy to retrieve, it can be used in other projects, and critical information like which specimen voucher is linked to a DNA extraction is not lost if people move on from RBGE.

Getting an EDNA number involves filling in a simple Excel spreadsheet with some basic collection information, and uploading it to a database. The Excel spreadsheet is accessible to RBGE lab users on an internal server (DNA, Molecular lab registration forms, EDNA (DNA), EDNA_submission_sheet_v01), and has two sets of fields, required and additional. If anything’s missing from the required fields, an EDNA number will not be issued, whereas the additional data is recommended but not essential… However, the more fully complete the data entry is, the faster it is to use it to generate GenBank submissions and publication voucher tables, justifying spending a little extra time on getting the forms completed.

Two points to remember when filling in the spreadsheet are not to use special characters, and not to make any of the entries too long, as there’s a maximum character number.

 

REQUIRED INFORMATION

Taxon name: this should not have authority information (Bellis perennis L.), just the genus and specific epithet (Bellis perennis).

Collector name: this cannot begin with an initial (J. Smith) as it will be rejected by the database; either use a full Christian name (John Smith), or put the surname first (Smith, J.).

Collector number: if there is none, s.n. is accepted.

Country code: two-letter standard codes; when filling in the spreadsheet, there is a tab with all the codes that you can look up (e.g. DE for Germany).

Material type: drop-down menu choices – fresh, frozen, herbarium, seed, silica gel dried.

Extraction type: drop-down menu choices include tissue maceration type, e.g. pestle, or mixer mill, and chemistry used, e.g. CTAB, Plant DNeasy minikits, Qiextractor.

 

ADDITIONAL INFORMATION

User DNA ID: this is the number that was given to the extraction in the lab; it’s extremely useful to have this for various troubleshooting in the lab – it can help match accessions to tubes, sort out issues with sample order, etc.

Extraction Date: entered in standard format year-month-date. Again, this can be useful for later troubleshooting, e.g. for separating batches of extractions by date, in case something went wrong on a particular day.

Herbarium barcode: this is ONLY for RBGE herbarium barcodes, not those from other institutes. If this is available, filling this in will propagate specimen data from the herbarium database. However, the required fields still need to be filled in.

Living Accession Number: this is ONLY for RBGE living accessions, not those from other institutes. If this is available, filling this in will propagate specimen data from the living collection database. However, the required fields still need to be filled in. The qualifier letter should not be filled in here.

Living Qualifier: this field is for any alphabetical character after the Living Accesion Number.

Silica Gel Box Number: this field is best left empty unless silica material came from a box numbered in the same format as “SGN12345”.

Sample note: free field, but there is a limit on how many characters are allowed, so should be kept short, and free from special characters. It may be useful to note e.g. if the extraction was from sporophyte versus gametophyte tissue, or flower versus leaf.

Location: free field, but there is a limit on how many characters are allowed, so should be kept short, and free from special characters.

Coordinates: free field, but there is a limit on how many characters are allowed, so should be kept short.

Decimal longitude:

Decimal latitude:

Collection Date Verbatum: this is for dates that cannot be turned into the correct date format, e.g. “Spring 1920”, “October 1976”.

Collection Date: entered in standard format year-month-date. This can be very useful in relation to DNA quality. If this is filled in, there is no point also filling in the Collection Date Verbatum field.

Note: free field, but there is a limit on how many characters are allowed, so should be kept short, and free from special characters.

 

Once the EDNA form is filled in, it can be uploaded to the EDNA database, which is available to users at RBGE who have a Username and Password.

Once logged on, the tab ‘Importer’ becomes highlighted; at the bottom of the Importer screen is a “Load” button.  The information in the excel sheet should be pasted into the ‘Load data’ window, and mapped to the fields. This will leave four fields that need to be filled in manually, three required fields: User (the lab user’s name, available from a drop-down list); Project (again, from a drop-down, e.g. MSc, barcoding, Leguminosae); Contact (a permanent staff member who will take long-term responsibility for the project, chosen from the drop-down list) – and one optional field, EDBANK Format (how the DNA will be stored long term – Plate, Strip or Tube; for most phylogenetics projects DNA will be stored in individual tubes, while for some population genetic project it will be stored in strips or plates – check with the molecular lab staff if unsure which format to chose).

After this information is filled in, the tab “Validate” becomes available. The entered data is screened for things like collector names that start with initials, accession numbers, dates, latitudes and longitudes that are in the wrong format, or other errors. If any are found, then these need corrected in the excel spreadsheet and the information all needs reloaded and re-entered. If there are no validation errors, the “Import to EDNA” button becomes available. At this point, the data will either successfully import, or other errors will be identified (e.g. non-standard characters, or too many characters). Unfortunately errors identified at this later point only stop EDNA numbers being generated for individual samples rather than for the whole batch, and it is not possible to cancel the issued EDNA numbers. This means that, for example, if entering a plate of 96 DNA extractions to EDNA, it’s quite possible for some samples in the middle of the plate to not be assigned a number. Obviously this becomes a sample labelling headache that is optimally sorted by redoing the entire batch to get consecutive EDNA numbers for all the samples, although this will lead to apparent duplicates of samples in the database. Molecular lab staff should be informed of redundant numbers, so that the duplicates are not also assigned places in the DNA bank.

When the numbers have been generated, they can be downloaded from the database by clicking on the “Tasks” tab, and the “As Spreadsheet” option – this will return all the information that has just been entered, along with the EDNA accession numbers for each sample.

 

See also:

The RBGE DNA bank

Jul 042016
 
Inga umbellifera Matthews

Fig. 1. Inga umbellifera, Mathews 1593, collected in Peru, Departamento San Martin: Provincia San Martin, Tarapoto, in 1835.

As part of our hybrid capture project, we sampled from an Inga umbellifera specimen that was collected about 180 years ago, by Andrew Mathews, in Peru in 1835. We made our Qiagen Plant DNeasy DNA extractions from the piece of plant tissue shown in Fig. 1. The DNA was very degraded, and present in quite low quantities (Tapestation fragment size distribution 46 to 306 bp; Qubit concentration 1.23 ng/μl; Fig. 2); from this we were still able to generate both Illumina Tru-Seq and NEB-Next DNA libraries (#13) that we sent to Edinburgh Genomics sequencing facility at the University of Edinburgh for Illumina Mi-Seq sequencing.

Inga umbellifera DNA from 1835 herbarium sheet

Fig. 2. Tapestation trace for Inga umbellifera DNA from Mathews’ 1835 herbarium sheet

We analysed the sequence data from the Tru-Seq and NEB-Next libraries separately, as our study aimed to compare different DNA extraction and library preparation techniques; from each library, we obtained over 300 thousand bases of Inga DNA sequence data that matched to our hybrid bait set. For the two libraries combined, we obtained over 1.6 million reads that passed quality controls, over 85% of which matched the hybrid baits that we’d used, and another c. 5% of which matched the Inga plastid genome. However, that still leaves around 10% of the reads that were not from the legume hybrid baits that we’d used, and we were interested to see what else might be present in that data set.

Library No. of trimmed reads % reads aligned to baits % reads aligned to Inga plastid Average quality score of variant positions (AQV) Number of variant bases Loci recovered (max 276) Conservatively called sequence (CCS), bp
H1835_NEB13+ 1,013,414 87.40% 4.30% 139.18 7,186 249 317,244
H1835_Tru13+ 659,161 84.20% 5.20% 132.97 7,045 247 310,949

 

We compared the DNA sequences of the reads from one of these libraries, H1835_NEB13+, to a publically available database (using a blast analysis run using Galaxy). This gave us a metagenomic analysis of the H1835_NEB13+ library reads, so that we could tell if anything other than Inga DNA had been in the extractions. Of 12,754,803 untrimmed reads, a huge number, over 8.3 million, were matches to legume sequences, exactly as we would expect from an Inga specimen (65%, with 1,211,319 Ingeae tribe reads, 2,015,461 Mimosioid legume reads, and 5,088,509 Fabaceae reads).

But not all the sequence reads were matches to plant DNA…

There were also:

3,047 reads that matched Homo sapiens (0.02%, which may or may not come from our intrepid plant collector, Mr Andrew Mathews, from his Peruvian wife, from the herbarium worker who stuck the pressed plant onto the card sheet, from taxonomists who have handled the specimen over the years, or indeed from anyone in the laboratory when the DNA extractions were being made…),

88,216 reads that matched Mediterannean mussels (0.7%),

and 346,820 reads that matched Streptococcus (2.7%; always wash your hands after touching herbarium specimens!).

Humans and bacteria are pretty easy to explain – people have been handing this plant material ever since it was collected, and there are bugs everywhere. But what about the mussels? The city of Tarapota in Peru is a long way from the coast, but after the plant was collected, it was squashed and dried in a plant press, and transported across the country, and eventually over to Europe. When we first got these results, we imagined working dinners in the Mathews household, with the botanist splitting his attention between his plant collections and a towering plateful of shellfish, dripping mollusc juices across the specimens. It does, however, seem unlikely that a professional plant collector would be quite that careless.

An alternative explanation is that we’ve also extracted DNA from glue. The leaf material we are working with has been removed from a herbarium sheet, to which it had been stuck. Animal-based glues were common in the 19th century, and although the classic glues were from mammal hides and fish, mussels certainly have a lot of sticky potential.

 

 

References:

Hart, M.L., L.L. Forrest, J.A. Nicholls & C.A. Kidner. In press. Retrival of hundreds of nuclear loci from herbarium specimens. Taxon.

James A. Nicholls, R. Toby Pennington, Erik J.M. Koenen, Colin E. Hughes, Jack Hearn, Lynsey Bunnefeld, Kyle G. Dexter, Graham N. Stone & Catherine A. Kidner. 2015. Using targeted enrichment of nuclear genes to increase phylogenetic resolution in the neotropical rain forest genus Inga (Leguminosae: Mimosoideae). Frontiers in Plant Science 6: 710. doi: 10.3389/fpls.2015.00710

 

Capturing Genes from Herbaria. I. What it’s all about. http://stories.rbge.org.uk/archives/16411

Capturing Genes from Herbaria. II. Inga. http://stories.rbge.org.uk/archives/16427

Capturing Genes from Herbaria. III. The samples. http://stories.rbge.org.uk/archives/16441

Capturing Genes from Herbaria. IV. DNA. http://stories.rbge.org.uk/archives/16470

Capturing Genes from Herbaria. V. Fragmenting the DNA. http://stories.rbge.org.uk/archives/16525

Capturing Genes from Herbaria. VI. Size Selection. http://stories.rbge.org.uk/archives/16645

Capturing Genes from Herbaria. VII. Comparisons. http://stories.rbge.org.uk/archives/16737

Capturing Genes from Herbaria. VIII. Amplification. http://stories.rbge.org.uk/archives/16788

Capturing Genes from Herbaria. IX. Hybrid capture. http://stories.rbge.org.uk/archives/17298

Capturing Genes from Herbaria. X. An update. http://stories.rbge.org.uk/archives/20751

Capturing Genes from Herbaria. XI. Some metagenomics of a herbarium specimen. http://stories.rbge.org.uk/archives/20817

Jun 302016
 

Last May (the 15th, to be precise), we sent three eppendorf tubes containing Illumina Tru-Seq and NEB-Next libraries constructed from Inga DNAs, most of which had been extracted from herbarium specimens, to the Edinburgh Genomics sequencing facility at the University of Edinburgh. A series of Botanics Stories blogs (listed below) described the rationale for the project, and the methodology that we followed in order to fill these tubes with libraries. Then followed a bit of a hiatus, so it seems that a quick update is in order.

In short: We got the data back on the 1st of June 2015, analysed it – it worked – and wrote a brief paper that was submitted to the journal Taxon on the 4th of February 2016, and accepted on the 19th of May. This was followed by a pleasant celebratory lunch in one of the local bars, the Orchard, on Wednesday the 25th… but back to the data:

Data from Hart et al. (Taxon, in press 2016): The results of the MiSeq runs, by library, are given in the following table. The final row is the silica-dried material from Dexter 401 (E) sequenced by Nicholls & al. (2015), for comparison with libraries from H2004. The library names start with H or S, depending on whether the DNA was extracted from herbarium specimens or silica-preserved tissue samples, followed by the collection year for each accession; the second part of each name reflects the library preparation kit (Tru-Seq or NEB-Next) and whether or not the DNA was repaired (+ or -), with a number that links back to previous blogs on DNA extraction, fragmentation, size selection and library preparation methods.

 

Library No. of trimmed reads % reads aligned to baits % reads aligned to Inga plastid Average quality score of variant positions (AQV) Number of variant bases Loci recovered (max 276) Conservatively called sequence (CCS), bp
H1835_NEB13+ 1013414 87.4% 4.3% 139.18 7186 249 317244
H1841_NEB07+ 214315 53.9% 0.7% 101.80 883 137 46045
H1841_NEB08+ 365550 73.2% 0.8% 73.44 2773 226 120148
H1932_NEB11+a 1226043 89.0% 4.7% 157.83 6377 248 322337
H1932_NEB11+b 862599 89.1% 4.1% 141.89 6253 246 310470
H1932_NEB11+bv2 1152606 90.0% 2.3% 133.15 5930 248 301994
H1932_NEB12+ 1919229 87.4% 6.4% 173.56 6463 249 331326
H1948_NEB05+ 583010 87.4% 1.6% 94.94 5028 239 241758
H1948_NEB06+ 704977 87.1% 3.7% 136.32 6132 247 298809
H2004_NEB09+ 1787314 74.3% 9.2% 168.53 7018 248 328618
H2004_NEB10+ 1595602 80.3% 10.4% 174.46 7135 250 334242
H2009_NEB01- 1711918 75.0% 8.6% 169.24 6482 248 326187
H2009_NEB01+ 1658799 76.6% 8.2% 169.21 6484 250 324340
H2009_NEB02- 1355984 75.2% 8.3% 163.79 6525 247 322957
H2009_NEB02+ 1668026 76.2% 8.5% 171.90 6516 250 326466
H2009_NEB03- 1513515 73.8% 8.3% 162.85 6463 246 319683
H2009_NEB03+ 1504758 74.0% 8.4% 161.80 6419 245 320273
H1835_Tru13+ 659161 84.2% 5.2% 132.97 7045 247 310949
H1932_Tru11+a 1584437 87.7% 3.8% 155.89 6246 248 322199
H1932_Tru11+b 1015706 87.5% 3.8% 144.88 6194 248 314862
H1932_Tru11+b2 1416246 87.0% 4.6% 159.42 6448 249 324910
H1932_Tru12+ 1774508 84.4% 6.3% 169.72 6503 248 330462
H1948_Tru05+ 1042441 83.9% 2.6% 136.01 5941 248 296844
H1948_Tru06+ 892927 84.6% 3.9% 145.22 6211 247 308853
H2004_Tru09+ 1958838 77.9% 9.2% 173.90 7041 249 333904
H2004_Tru10+ 1576572 77.4% 9.5% 170.05 7066 248 330278
H2009_Tru01- 1338317 77.6% 9.1% 167.51 6601 249 324201
H2009_Tru01+ 1536759 77.2% 8.4% 167.03 6594 248 325184
H2009_Tru02- 1476338 76.6% 8.4% 166.63 6569 249 323881
H2009_Tru02+ 1226123 75.6% 8.5% 161.46 6572 249 319045
H2009_Tru03- 1630041 75.4% 8.6% 168.09 6509 250 324451
H2009_Tru03+ 1753019 75.0% 8.4% 167.90 6512 249 323951
S2004_TruKD401 689439 74.4% 9.2% 156.29 5809 245 330396

 

References:

Hart, M.L., L.L. Forrest, J.A. Nicholls & C.A. Kidner. In press. Retrival of hundreds of nuclear loci from herbarium specimens. Taxon.

James A. Nicholls, R. Toby Pennington, Erik J.M. Koenen, Colin E. Hughes, Jack Hearn, Lynsey Bunnefeld, Kyle G. Dexter, Graham N. Stone & Catherine A. Kidner. 2015. Using targeted enrichment of nuclear genes to increase phylogenetic resolution in the neotropical rain forest genus Inga (Leguminosae: Mimosoideae). Frontiers in Plant Science 6: 710. doi: 10.3389/fpls.2015.00710

 

Capturing Genes from Herbaria. I. What it’s all about. http://stories.rbge.org.uk/archives/16411

Capturing Genes from Herbaria. II. Inga. http://stories.rbge.org.uk/archives/16427

Capturing Genes from Herbaria. III. The samples. http://stories.rbge.org.uk/archives/16441

Capturing Genes from Herbaria. IV. DNA. http://stories.rbge.org.uk/archives/16470

Capturing Genes from Herbaria. V. Fragmenting the DNA. http://stories.rbge.org.uk/archives/16525

Capturing Genes from Herbaria. VI. Size Selection. http://stories.rbge.org.uk/archives/16645

Capturing Genes from Herbaria. VII. Comparisons. http://stories.rbge.org.uk/archives/16737

Capturing Genes from Herbaria. VIII. Amplification. http://stories.rbge.org.uk/archives/16788

Capturing Genes from Herbaria. IX. Hybrid capture. http://stories.rbge.org.uk/archives/17298

Capturing Genes from Herbaria. X. An update. http://stories.rbge.org.uk/archives/20751

Capturing Genes from Herbaria. XI. Some metagenomics of a herbarium specimen. http://stories.rbge.org.uk/archives/20817

Apr 212016
 

What do Giant panda eat?  The answer might seem obvious but the reality is far from simplistic.

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One of Edinburgh Zoo’s Giant pandas Yáng Guāng eating bamboo

The diet of the Giant panda (Ailuropoda melanoleuca) is highly specialised on bamboo, but there are hundreds of different bamboo species and over 60 of them are eaten by Giant panda. The bamboo species consumed vary geographically and seasonally and even individuals appear to differ in their choices. All these different bamboos vary in their nutritional content and digestibility and Giant pandas can be very picky about which bamboo they eat and when. To further complicate things panda may also feed on other plants, fungi and even other animals, and we have no information on the importance of these non-bamboo foods to panda health and breeding.

Researchers have been investigating pandas for decades and we do know a lot about them but the finer details of the diet have eluded us – so far.

There are two key difficulties in assessing panda diet.

  1.  the full range of items consumed is difficult to determine from watching panda due to their reclusive nature
  2.  bamboo species are very difficult to tell apart, especially once eaten and digested by a panda.

But we now have tools to answer this complicated question

Scats from giant panda will be collected and tested to work out what the panda has eaten

Scats from Giant panda will be collected and tested to work out what the panda has eaten

Using DNA-based methods to investigate diet, we don’t have to find pandas, only their scats (their poo). We can extract the DNA from the scat, sequence it and then compare the scat DNA sequences to the DNA sequences of known bamboo species, or other plants, fungi or animals to work out what was eaten. However, this process is particuarly complicated in bamboo because they are difficult to tell apart even using DNA. So first we have to develop methods to focus on the parts of the DNA sequence that will allow us to distinguish between the different bamboo species. Once we can accurately tell them apart we will be able to find out a lot more about bamboo and the species that eat them.

In stage 1, Yáng Guāng (meaning “sunshine”) and Tián Tián (meaning “sweetie”), Edinburgh Zoo’s two giant pandas will be helping to develop these methods by providing samples to test these new methods on. This is an important step. We know exactly what has been fed to these captive pandas so we can determine how accurate and reliable our methods are.  Then we will move onto stage 2 – using these methods to investigate diet in wild panda.

This project is an international collaboration between RBGE and several organisations including the Leverhulme Trust, Royal Zoological Society of Scotland (RZSS), the Kunming Institute of Botany and the Institute of Zoology both at the Chinese Academy of Science, China Conservation and Research Centre for Giant Pandas, Panda Centre within the Wolong Nature Reserve, Edinburgh Consortium for Giant Panda Conservation and Forest Landscape Restoration and the Australian Museum Research Institute and we adding to the list all the time!

We are only just starting this exciting project but it has already attracted a lot of attention. Check out the short segment from CBBC Newsround!

Follow our work on Giant panda diet on Botanics stories where there will be regular blogs about the project and fieldwork (and on twitter @Linda_Neaves)!

Jan 302016
 

This last week I’ve actually managed to spend a bit of time in the lab, trying to get some gaps filled in a DNA barcoding matrix for simple thalloid liverwort Aneura. David Long and I are heading off to Trondheim in just over a week to combine our data set with one generated by Ana Maria Séneca Cardoso, working with Lars Söderström and Kristian Hassel at NTNU.

A fridge-full of DNA at RBGE

A fridge shelf piled with racks and plates of DNA in the RBGE PCR lab

Many of the DNA extractions that I have been trying to amplify are old (with a very few that were extracted 14 years ago at Southern Illinois University). Most of them have already been tried, and have previously failed to amplify, for the four selected barcode markers (three plastid genes, rbcL, rpoC1 and matK, and one plastid intergenic spacer region, psbA-trnH; a nuclear marker, ITS2, was originally included, but proved difficult to get good sequence data from). Most of the DNA extractions I’ve needed were scattered across a large number of Edinburgh DNA bank (EDNA) plates, while some of the rest of it had never been aliquoted out of the original QiaXtractor plates.

Our hard-working PCR machines wait for samples in the molecular lab

Our hard-working PCR machines wait for samples in the RBGE molecular lab

Because the amount of time needed to chase down the DNA samples was more than the amount of time needed to set up the reactions, and because my default protocol has changed, over the course of this project, from using CES as a PCR enhancer to using TBT-PAR as a PCR enhancer, I decided for three of the loci to test the amplification with both CES and TBT-PAR, building on from a previous Botanics Stories posting. (The exception was matK, for which we use a different polymerase, Invitrogen’s Platinum, and 5M betaine as a PCR enhancer; we have found that this gives better sequence reads than amplification with a standard Bioline taq does.)

getting ready for PCR - reagents defrosting on ice

Getting ready for PCR – my reagents defrosting on ice

Reactions were set up in 20 ul, using exactly the same reagents (Sigma water, 5x buffer, magnesium, dNPTs and forward and reverse primers, with 1 ul each of the Aneura DNA extractions), with the exception of the 4 ul of either CES, or TBT-PAR, per sample.

Inside the laminar flow hood and ready to go

Inside the laminar flow hood and ready to go

The PCR products were all run out on standard 1% agarose TBE gels, at 80 volts, for 40 minutes, stained with SybrSafe, and visualised under a blue light, to test for amplification success. As a size standard, 3.5 ul of an Invitrogen 1 kb ladder was loaded at intervals on each gel.

Several of these gels are shown below. In these images, DNA is stained so that it fluoresces in bright blue or UV light. The samples have been loaded into the gel in holes, or wells, that can be seen at the top of, and at regular intervals down, the gel, and migrate through an electric current towards the positive electrode that would be at the bottom of the image. Smaller fragments move faster through the gel, meaning that samples can be separated by the lengths of the DNA fragments.

The comparisons of amplification success with CES and TBT-PAR are unfortunately ambiguous.

For the psbA-trnH region, using TBT-PAR was more successful than using CES – as can be seen particularly clearly in the second row down, where only 3 of the 8 extractions amplified with the CES additive, but all 8 amplified with TBT-PAR.

Aneura DNA amplified for psbA-trnH region: left hand side - with CES additive; right hand side - with TBT-PAR additive

Aneura DNA amplified for the psbA-trnH region: left hand side – with CES additive; right hand side – with TBT-PAR additive

For both rbcL and rpoC1, it’s harder to get a clear picture. Some samples amplified with one additive rather than the other, while most samples amplified with both.

Aneura DNA amplified for rpoC1 region - first two rows with TBT-PAR additive; second two rows with CES additive

Aneura DNA amplified for rpoC1 region – first two rows with TBT-PAR additive; second two rows with CES additive

For a set of the rpoC1 amplifications, although all samples amplified using both additives, the bands from the reactions with CES (the three lowest rows in the gel image) are rather brighter than those from the reactions with TBT-PAR (the upper three rows), meaning that there is more amplified product in them. However, the CES bands do appear slightly more smeary, so it may be worth comparing the quality of sequence data from both sets of reactions as well as just considering amplification success.

The gels for the rbcL samples (shown below) are harder to interpret – the consistent bright bands are not the PCR product that I am looking for, but represent an artefact of the reaction known as “primer dimer”; the PCR product is the second slower (and therefore, longer) DNA fragment that is sometimes present. There seem to be more samples that have amplified with the CES additive, although a few samples that have failed with CES have instead amplified with TBT-PAR, meaning that in this instance, having used both protocols in parallel will allow me to generate DNA sequences from more accessions of Aneura than I would have been able to, had I just used one PCR protocol.

Aneura DNA amplified for rbcL plant barcode region with CES additive

Aneura DNA amplified for rbcL plant barcode region with CES additive

Aneura DNA amplified for rbcL plant barcode region with TBT-PAR additive

Aneura DNA amplified for rbcL plant barcode region with TBT-PAR additive (2 gel images)

160128 Aneura rbcL M745 TBT 7to9 crop

The only recommendation that I can think of from this is that, if time is pressing, the DNA samples are difficult to access, and you need as much amplification as possible as fast as possible, do two sets of PCR reactions, using both additives.

This does, however, have the unfortunate side effect of doubling the cost of your PCR, as well as giving you twice as many samples to load onto your gels…

 

See also: Botanics Stories: Sparking additions in the Molecular Lab. http://stories.rbge.org.uk/archives/2271

Nov 032015
 

Distributions of Delongia cavallii (circles), which occurs in the East African Rift Mountains and on Réunion, and D. glacialis (squares) which spans the Himalaya from Pakistan to Yunnan

The relative structural simplicity of some groups of mosses can disguise their uniqueness, especially when simplified features have evolved multiple times within the same family from ancestors with more complex morphologies. The family Polytrichaceae is particularly well-known for very robust mosses such as the familiar Polytrichum commune, in which the leaves have sheathing bases and densely packed longitudinal “walls” of cells (lamellae) on their upper surfaces to increase the area available for the uptake of carbon dioxide and so the rate of photosynthesis in the exposed habitats they tend to grow in. This “pseudomesophyll” gives them an opaque appearance superficially similar to the leaves of vascular plants (despite the very different structure), and unlike the translucent, single-cell-layered leaves of most mosses. However, within the family a number of lineages have reverted to a smaller, more typically moss-like growth form with simpler leaves and much reduced lamellae, and it can often be difficult to know whether these share a common ancestor with such features or have developed them independently.

David Long on fieldwork in Nepal (photo by David Knott)

Piton des Neiges on Réunion, where Delongia cavallii (then Oligotrichum cavallii) was found in 2011, considerably extending its range beyond mainland East Africa (photo by Terry Hedderson)

Slopes above Yume Samdong in North-east Sikkim, a locality for Delongia glacialis. The large plant in the foreground is Rheum nobile, the Sikkim rhubarb (photo by David Long)

We have known for a few years that the smaller, morphologically simplified Polytrichaceae found in southern temperate parts of the world are not at all closely related to the similar looking species in Asian, north temperate and arctic regions placed in the genera Oligotrichum and Psilopilum. But even within Asia and the tropics it appears that there were two or three independent origins of this type of plant. In a paper just published in the journal TAXON we describe a new genus, Delongia, in honour of the recently officially retired (but still very active!) RBGE bryologist David Long, which includes two species previously placed in Oligotrichum. The name is particularly appropriate as most of the known collections of one of the species, Delongia glacialis, were made by David during his expeditions to Nepal, Sikkim and Yunnan and were first identified as Oligotrichum glaciale by Isuru Kariyawasam, an MSc student at RBGE. Interestingly, the second species, Delongia cavallii, has a quite different distribution, occurring in the East African Rift mountains and recently discovered by Terry Hedderson on the Island of Réunion.

Although molecular data were instrumental in recognising the distinctness of the new genus and its relatively distant relationship to Oligotrichum, it is also united by a number of unique anatomical features, not least spore capsules with a peculiar “spongy” lower part in which the stomata are often hidden when the capsule is dry and exposed when wet. In fact, this spongy texture had been noticed seventy years ago in O. cavallii and the rather euphonic generic name “Spoggodera” (“sponge neck” in Greek!) proposed, although this was never validly published.

A spore capsule of Delongia glacialis showing the "spongy" neck. Although the stomata are fairly exposed in this picture they become much more sunken when the capsule is dry

A spore capsule of Delongia glacialis showing the “spongy” neck. Although the stomata are fairly exposed in this picture they become sunken in distinct pits when the capsule is dry

We were able to use a type of “molecular clock” dating technique to estimate when the lineages of the two fairly different species of Delongia became separated from each other, as well as when Delongia separated from its most closely related extant genus (either Psilopilum or Atrichum s.l., not Oligotrichum). It seems that the most likely date for the separation of the Himalayan D. glacialis from the East African D. cavallii was about 23 million years ago, right at the boundary between the Oligocene and Miocene epochs and just at the time that the East African rift system was beginning to form (the Himalaya were also continuing to rise rapidly at this time).

The lineage of Delongia itself probably originated way back in the Eocene (56–34 million years ago), highlighting the importance of recognising and naming such unique and relatively evolutionarily isolated components of biological diversity that might otherwise be mistaken for recently evolved species with many close relatives. So although Delongia only has two species it certainly deserves its generic status. The value of taxonomy is that while experts might not need names to recognise the evolutionary diversity in their own specialist groups, it is only by representing this diversity in a universal nomenclature that it becomes generally accessible and quantifiable.

A chronogram (a tree with branch lengths representing absolute time) showing the likely origin of Delongia in the Eocene and the separation of the lineages of the two species around the Oligocene Miocene boundary

Bell, N.E., Kariyawasam, I., Hedderson, T.A. & Hyvönen, J. 2015. Delongia gen. nov., a new genus of Polytrichaceae (Bryophyta) with two disjunct species in East Africa and the Himalaya. Taxon 64: 893-910.

Oct 302015
 
Archegoniophores and antheridiophores of Marchantia; taken by Julia Bechteler

Archegoniophores and antheridiophores of Marchantia; photograph by Julia Bechteler

One of the earliest plastid genomes to be sequenced, in the late 1980s (Ohyama et al.), was that of Marchantia polymorpha, one of the commonest liverworts around town, and an increasingly widely used model organism for genetic research. The complete mitochondrial genome followed, with a key publication by Oda et al. in 1992. The research on both organellar genomes came from Kyoto University, Japan.

When we were generating a DNA sequence matrix for the complex thalloids, we also included GenBank sequences from the published genome sequences for both these organelles (mitochondrial loci nad1, nad5 and rps3 and plastid loci atpB, cpITS, psbA, psbT-psbH, rpoC1, rbcL and rps4), to see how they compared with sequences from the three subspecies of Marchantia polymorpha that are found in the United Kingdom (subspecies ruderalis is the weedy plant that is commonly found in paving cracks, flowerbeds and plant pots).

Marchantia phylogeny based on Villarreal et al. 2015, Fig. 2

Marchantia phylogeny based on Villarreal et al. 2015, Fig. 2

The results of the analyses (Villarreal et al. 2015) were rather unexpected: the plant from GenBank didn’t cluster with our Marchantia polymorpha accessions, but instead with a Marchantia paleacea accession from Mexico. Admittedly, both species form a clade, but there’s a convincing amount of genetic distance between them.

Marchantia polymorpha ventral scales, photograph by Des Callaghan Licensed under CC BY-SA 4.0 via Wikimedia Commons - https://commons.wikimedia.org/wiki/File:Marchantia_polymorpha_scales.jpg#/media/File:Marchantia_polymorpha_scales.jpg

Marchantia polymorpha ventral scales; photograph by Des Callaghan. Licensed under CC BY-SA 4.0 via Wikimedia Commons

 

Helene Bischler-Causse (1989, 1993) separated Marchantia into sections based on morphological characters of the plants, placing Marchantia polymorpha in section Marchantia, and Marchantia paleacea alone in section Paleaceae, due in part to differences in the way the scales on the underside of the thallus are arranged, as well as spore morphology. Now it seems that the organelle genome sequences that have, for the last 1/4 century, been thought to be from Marchantia polymorpha, are in fact from this lesser known (but also widespread) species, Marchantia paleacea.

(Kijak et al.’s research at Mickiewicz University in Poland shows a similar picture; although their study is still unpublished, a poster from the group is available to download here.)

New Phytol title