Analysing phenotypes and measuring callose

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Published on: January 9, 2013
Part of Figure 7 from Green et al., showing the an example of Phenophyte output.

At the end of last year, you may have missed two useful publications from Plant Methods which use new free online tools to make your life easier.

Phenophyte can help you measure 2D areas quickly and accurately. It was described in November’s Plant Methods by Green et al., a team mainly from  Columbia, USA. Users chose if they want to analyse indivudial images, compare before/after images (as shown in the figure to the left), or analyse a timecourse. They then upload the images – the upload tool allows up to 2GB or 500 images, and sequential uploads are possible if required. The computational results can be previewed before submitting the job. When processing is complete, the user will be emailed a link to the results, which must be downloaded within a week. The manual provides detailed tips on how to take the photographs to upload, and the guidance is standard with the exception of the use of a colour/size checker (for example, this one), and the interface is straightforward and friendly.

Figure 5 from Zhou et al., showing the CalloseMeasurer interface and output.

A more specialized application is CalloseMeasurer, from the Robatzek group at The Sainsbury Laboratory. Zhou et al. describe a piece of software for quantifying callose deposition with enough accuracy to quantify the growth of filamentous pathogens within a plant by recognising the spreading network of callose deposition caused by the pathogen. The paper is heavy on technical detail, but guides readers through using CalloseMeasurer in the ‘Image Processing’ section of the paper. Users must have Acapella software installed, and they simply drag and drop the CalloseMeasurer script into the application window and start using the programme.

How many ways can you measure a plant?

Categories: GARNet, guest blogger, methods
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Published on: January 8, 2013

In December, Ruth gave a talk at the Julich Plant Phenotyping Centre – here she explains what’s going on in plant phenotyping at the moment. 

Recently I had the opportunity to visit and talk at the Julich Plant Phenotyping Centre in Germany and see the wealth of tools and technologies that the centre has available to measure and analyse plant growth and development in a non invasive manner. By using a range of sensors and computer vision tools for quantifying plant traits the centre aims to help overcome the current bottleneck in effectively linking genotype to phenotype.

As a mere amateur in this field, I used CCD cameras during my Ph.D to monitor circadian rhythms and during my post-docs I just counted leaves to determine flowering time. I was amazed by the depth and breadth of analysis that can now be carried out, and on such a large scale.

For example their purpose built automated Rhizo screen enables researchers to non-invasively obtain quantitative measurements of root architectures of plants grown in soil in 2D as well as evaluating shoot area. Whilst a variety of spectral and optical imaging systems sensitive to a wide range of wavelengths provide a plethora information from chlorophyll fluorescence, water content, lignin and cellulose composition to growth dynamics via leaf area. The centre also has a NMR, MRI and PET setup to visualize the inner structure of plant organs and tissue and transport of substances such as CO2. (Fiorani et al. Imaging plants dynamics in heterogenic environments. Current Opinion in Biotechnology, 23: 227-235).

Julich is just one of a number of phenotyping centres that are being established all over Europe, including the UK centre at Aberystwyth. The major European centres have been linked together in the European Plant Phenotyping Network (EPPN). This network offers access to 23 different plant phenotpying facilities spread across the EU. So if you haven’t experienced the power of phenomics yet this might be one way to dip your toe in phenotyping water!

Imaging trichomes

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Published on: December 13, 2012
cryo-scanning electron microscope image of a trichome on an Arabidopsis leaf

Even if you don’t work on trichomes, you have definitely experienced them first-hand, as stings on nettles are trichomes that have evolved down a particularly nasty route. Other trichomes pack less of a punch, but they are still important for phytochemical production and herbivore defence.

Arabidopsis trichomes are rather more tame than stinging nettle trichomes, and present an excellent way to study cell differentiation as well as being a target for crop improvement. But although trichomes are easy to see using light microscopy, they are difficult to study. Manually counting them and recording their length and position is tiresome in the extreme, and imaging technologies tend to require specialized skills and software that not all labs can access.

In today’s highlighted paper, Pomeranz et al. present a new method of analysing trichomes consisting of polarizing light microscopy (PLM) and a web-based imaging tool. In fact PLM is an old technique described by Ballard in 1916, and is an excellent way of imaging trichomes because of the highly crystalline cellulose in trichome cell walls which confers polarizing (birefringent) properties. As the authors say, this new technique is a ‘repurposed’ method, and the key to the novel technique is the online resource TRICHOMENET, which allows imaging and easy analysis of trichomes, and can be linked with ImageJ.

It certainly appears that this method would be easy to set up in any lab. Preparing samples for PLM involves methanol or ethanol, lactic acid, and a water bath – the method is in the paper or in Bischoff et al. (2010). PLM itself requires polarizing filters, which can be bought in a kit, for example from Motic, or as individual filters. The image is then uploaded to TRICHOMENET, which guides the user through counting the trichomes. Once the data is recorded, TRICHOMENET can analyse trichome positional data, density, and distances.

Highlighted article: Marcelo Pomeranz, Jeffrey Campbell, Dan Siegal-Gaskins, Jacob Engelmeier, Tyler Wilson, Virginia Fernandez Jelena Brkljacic, and Erich Grotewold (2012) High-resolution computational imaging of leaf hair patterning using polarized light microscopy. The Plant Journal ‘Accepted Article’, doi: 10.1111/tpj.12075

Image credit: Emmanuel Boudet.

Progress in pollen research

Time lapse video of Arabisopsis pollen grains germinating and growing pollen tubes. Credit: .

As an officer for GARNet, the Arabidopsis research network, I am happy to share the news that we can now add pollen germination to the long list of things for which our Arabidopsis can be called a model plant. The research is published in New Phytologist, and is currently in early view.

The importance of studying pollen for plant reproduction research is obvious, but it is also an excellent and widely used system for studying cell growth and development. Some plants, such as tobacco, have pollen that can be germinated on cue, and monitored in all sorts of ways as through germination, cell development, and pollen tube growth. Unfortunately brassicas, including Arabidopsis thaliana, do not have such amenable pollen.

A team of researchers from Oxford have developed a method that yields fast, reliable germination of A.thaliana pollen. The pollen tubes that grow are long and morphologically normal.

The method uses a cellulose-based membrane covering an agarose pad, all set up on a glass microscope slide. In the authors’ view, this protocol was more successful than other attempts because the environment surrounding the pollen mimics the stigma – so not only does this paper present a method of studying Arabidopsis pollen, but it provides novel information about the environmental cues required for pollen germination. The method was optimized for temperature and pH as well as the ratios of reagents used to make the materials.

Although this paper was about Arabidopsis and marks an important development for Arabidopsis researchers working on pollen and cell growth, it is also significant for Brassica researchers. The Brassica family contains many commercially important crops, and this method can surely be adapted to serve research into cabbage, oilseed rape, or other Brassica species.

Highlighted article: M. J. Rodriguez-Enriquez, S. Mehdi, H. G. Dickinson and R. T. Grant-Downton (2012) A novel method for efficient in vitro germination and tube growth of Arabidopsis thaliana pollen. New Phytologist (Early View) doi: 10.1111/nph.12037


Categories: methods, resource
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Published on: November 6, 2012
Fig. 4 from Nieuwland et al. (2012), showing the Phytotracker labels

If you have ever been frustrated by a less than clearly labeled seed stock, not knowing what the green and yellow dots mean, how long its been in that drawer, or which generation it is, GARNet Chair Jim Murray’s lab in Cardiff have worked out a system that will help.

Phytotracker is a system that organizes your seeds for you. Of course it depends on people recording the tray number and parent lines in the database, and correctly labeling the seed stocks and plasmids in the lab. However, once you’ve done that, you can forget about it because Phytotracker does the remembering for you – everything from which plasmid was used for the transformation to when to harvest the seeds.

The system is well explained in the paper, which was published in Plant Methods in October. If you want to try the system out, you’ll need Filemaker Pro (version 8 or later), or for a fully networked solution Filemaker Pro Advanced (version 8 or later: currently Filemaker Pro is version 12). Your University may already have a site licence! You’ll also need printers in your growth rooms and labs to print labels for the trays, plants, and seed stocks. Commitment from everyone in your group is essential – this system would fall apart if you have a regenade group member who insists on labeling with autoclave tape and a Sharpie. It has been successfully used in Cardiff for five years though, so it looks like a system that is worth committing to.

Highlighted article: Jeroen Nieuwland, Emily Sornay, Angela Marchbank, Barend HJ de Graaf, James AH Murray (2012) Phytotracker, an information management system for easy recording and tracking of plants, seeds and plasmids. Plant Methods 8:43

Download Phytotracker here:


Friday Film: Automatic cell counting with ImageJ

Categories: methods, resource
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Published on: October 5, 2012

This is a video tutorial on quantifying cells using ImageJ. ImageJ is a free tool for image processing and analysis in Java. I chose to highlight this tutorial as it is clearly explained and may be of use to many plant scientists, but YouTube is a goldmine of other less professional tutorials on using ImageJ for any number of applications. I was particularly interested in this one on quantifying stained liver tissue, as I used to work on secondary cell walls and it would have been a handy tool for qualitative analysis of my many images of phloroglucinol stained tobacco stem cross-sections.

Created by Keene State College’s Center for Engagement, Learning, and Teaching.

Working with Natural Antisense Transcripts

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Published on: September 27, 2012

In January of this year Nucleic Acids Research published a paper describing a database of plant natural antisense transcripts (NATs), PlantNATsDB (Chen, Yuan et al., 2012). It contains around 2 million NATs from 69 plant species, and has a simple viewer showing the two loci involved in each NAT, any overlap, the NAT type, and an option for more detail. It is also possible for search for NATs for specific loci. It is important to note however that the database was last updated a year ago, in September 2011.

This month’s Plant Methods also features a NATs tool, a protocol for NAT identification in plant tissue (Collani and Baraccia, 2012).  It is a simple PCR based method, and relies on prior knowledge of the existence of a NAT, as specific primers are needed. Used in association with PlantNATsDB, this is a useful technique. (more…)

High-throughput, cheap, reliable DNA extraction method

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Published on: September 11, 2012

Highlighted article: Zhanguo Xin and Junping Chen (2012) A high throughput DNA extraction method with high yield and quality. Plant Methods 2012, 8:26 doi:10.1186/1746-4811-8-26

Judging by the fact that it was accessed 1400 times in less than a month, the DNA extraction method described by Xin and Chen in last month’s Plant Methods must be worth a look.

Having had a look, I can tell you it seems to be an invaluable method for cheap, reliable high-throughput DNA extraction. It works on seeds and leaves from a number of plants, and according to the abstract, one person can manage 192 extractions in a working day. Using the price estimation in the paper, this would cost about £13 in total for consumables.

The protocol is clear and easy to follow, setting out exactly what reagents, consumables and equipment you will need so there will be no panicked begging of microtitr plates from a friend halfway through the extraction. As well as standard lab equipment which should be accessible to most researchers, you will need MagAttract Suspension G. MagAttract provides  the simple, efficient clean up step, while the rest of the protocol is based on a traditional CTAB extraction method.

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