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SENSOIL: new generation of transparent soils for the study of rhizosphere processes


Figure 1: The thin layer of soil surrounding plant roots (Rhizospehre) is the site of various biological, physical and chemical processes

It has been estimated that about 70% of N-fertiliser applied worldwide is lost to the environment, leading to economic losses, water pollution and increased greenhouse gas (GHG) emissions. When nitrogen is in the form of nitrate (NO3-), it is relatively mobile in soil and thus can easily be acquired by plants over distance through mass flow. But nitrate is also subject to leaching and denitrification which, if incomplete, produces nitrous oxide emission, a gas with ca 300× the GHG effect of CO2. Ammonium (NH4+) is far less mobile in soil as it forms bonds with the soil particles so that losses by leaching are extremely low. Ammonium metabolism is also more favourable to plants since it requires 4 times less energy for assimilation. Large amounts of N-fertilisers are therefore supplied in the form of ammonium. A major agronomic concern, however, is the nitrification of ammonium fertilisers by soil microorganisms where large amounts of nitrogen are transformed to nitrate and thus reduce soil pH, and become highly mobile and susceptible to leaching and conversion to GHG (N2O) via denitrification. 

Plant and microbial processes in soil essential to the nitrogen cycle in soil. In particular, the thin layer of soil surrounding plant roots (rhizosphere, Figure 1) is the site of complex and intense biological activity.  Plants roots exude a wide variety of compounds in the neighbouring soil environment, and these exudates are sensed and exploited by a variety of soil micro-organisms. It has been shown for example that plant release chemical compounds that inhibit the growth of microbes involved in the transformation of nitrogen in soil. 

Live imaging of soils

Biological activity in the rhizosphere is highly dynamic. The nature, location and intensity of root secretions depend largely on the maturity of the root tissue, the anatomy of the root and the physiological status of the plant. Therefore, in order to understand how biological organisms establish and maintain in the rhizosphere, we must find ways to observe and measure soil biological activity live and in situ. This has proved extremely difficult using existing imaging technologies used in the natural sciences. Figure 2: Arabidopsis root in transparent soil imaged at high resolutionSoils are opaque and techniques available for analysis are often destructive for example, FISH and thin sectioning.

Penetrating radiation such as X-rays, neutron imaging, or magnetic resonance can be used to image roots, soil particles or water contents, but not microbial activity. Methods based on light imaging techniques provide the greatest insights into the functioning of soils because they can rely on a range of powerful microscopy techniques, fluorescent reporters and biochemical sensors. The main challenge, therefore, is to develop systems to observe soil biological organisms using microscopy and light imaging. 

Transparent soils

Recent research carried out by the team demonstrates that the physical and chemical structure of soils can be replicated using low refractive index (RI) solid particles that can adsorb nutrients and dyes and are suitable for image living organisms in situ. Our first generation of Transparent Soil (TS) exhibits physical properties and growth conditions similar to those in sandy soils. Water retention can exceed that of sand (similar to vermiculite) and particle surface charge density is high, which is suitable to model clay particles (1,2). Mixtures of particles containing a range of size, surface charge density and compaction level could be used to mimic a natural soil. TS offers new opportunities to soil biology because they allow roots, microbes and soil particles to be imaged at resolutions not previously achievable. 


They  are amenable to modern microscopy techniques, for example, Laser Scanning Microscopy, Optical Projection Tomography or BioSpeckle Laser imaging (3-5). They can host numerous fluorescent markers and other dyes so that complex biological and chemical activity can be quantified non-destructively. Quantitative analysis of image data based on fluorescence imaging is greatly facilitated and provide high contrast images that are suitable for computational processing of  data (Figure 2 and 3).

The project

The recent development of a transparent soil in our laboratories offers great scope to explore and unravel complex soil biological processes. The discovery of this new material have attracted much attention from the scientific community, and as part of this success, we are now looking to recruit enthusiastic scientists to study nitrification activity in the rhizosphere. This work is part of a prestigious award from the European Research Council (ERC) and involves multidisciplinary and international collaborations.

The broad aims of this project are therefore: 1) Engineer smart transparent soil technologies combining polymer chemistry, and soil physics; 2) design new instrument for live imaging of biological and chemical processes in soil 3) to quantify nitrogen movement and transformation in soil; and 4) to discover how the rhizosphere is formed, how microbial activity, soil structure and root activity influence nitrogen movement in soil. 

Research team

The project builds on a multidisciplinary team of scientists based at the James Hutton Institute in Dundee, the University of Dundee on and French CNRS. The team has expertise in computational and theoretical biology (Dr Dupuy), fluopolymers (Dr Ameduri and Ladmiral) and photonics (Dr MacDonald), soil chemistry and biology (Drs Daniell, Holden, Lumsdon), rhizosphere and plant nutrition sciences (Drs White and Geroge).

Further reading

  1. 1. Downie, H., Holden, N., Otten, W., Spiers, A.J., Valentine, T.A. and Dupuy, L.X. 2012. Transparent soil for imaging the rhizosphere. PloS one 7(9): e44276.
  2. 2. Downie, H., Valentine, T.A., Otten, W., Spiers, A.J. and Dupuy, L.X. 2014. Transparent soil microcosms allow 3D spatial quantification of soil microbiological processes in vivo. Plant Signalling and Behavior: e29878.
  3. 3. Yang, Z., Downie, H., Rozbicki, E., Dupuy, L.X. and MacDonald, M.P. 2013. Light Sheet Tomography (LST) for in situ imaging of plant roots. Optics Express 21(14), 16239-16247.
  4. 4. Braga, R.A., Dupuy, L.X., Pasqual, M. and Cardoso, R.R. 2009. Live biospeckle laser imaging of root tissues. European Biophysics Journal 38, 679-686.
  5. 5. Ribeiro, K.M., Barreto, B., Pasqual, M., White, P.J., Braga, R.A. and Dupuy, L.X. 2014. Continuous, high-resolution biospeckle imaging reveals a discrete zone of activity at the root apex that responds to contact with obstacles. Annals of Botany 113(3), 555-563.
  6. 6. Federici, F., Dupuy, L.X., Laplaze, L., Heisler, M. and Haseloff, J. 2012. Integrated genetic and computation methods for in planta cytometry. Nature Methods 9(5), 483-485.
  7. 7. Dupuy, L.X., Mackenzie, J. and Haseloff, J. 2010. Regulation of plant cell division in a simple morphogenetic system. Proceedings of the National Academy of Sciences 107 (6), 2711-2716. 
  8. 8. Duarte, I.M., Almeida, M.T.M., Duarte, M.M., Brown, D.J.F. and Neilson, R. 2011. Molecular diagnosis of trichodorid species from Portugal. Plant Pathology 60, 86-594.
  9. 9. Donn, S., Neilson, R., Griffiths, B.S. and Daniell, T.J. 2012. A novel molecular approach for rapid assessment of soil nematode assemblages - variation, validation and potential applications. Methods in Ecology and Evolution 3, 12-23.
  10. 10. Chen, X.Y., Daniell, T.J., Neilson, R., O’Flaherty, V. and Griffiths, B.S. 2014. Microbial and microfaunal communities in phosphorus limited, grazed grassland change composition but maintain homeostatic nutrient stoichiometry. Soil Biology and Biochemistry 75, 94-101.


Areas of Interest

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The James Hutton Research Institute is the result of the merger in April 2011 of MLURI and SCRI. This merger formed a new powerhouse for research into food, land use, and climate change.