RGB e Imaging spettrale radice per pianta Phenotyping e ricerca fisiologica: messa a punto sperimentale e protocolli di Imaging

Riepilogo

Un protocollo sperimentale è presentato per la valutazione del terreno coltivato pianta radice sistemi con RGB e l'imaging iperspettrale. Combinazione di tempo di immagine RGB serie con chemiometrica informazioni da iperspettrale scansioni ottimizza intuizioni nelle dinamiche di radice di pianta.

Abstract

Migliore comprensione delle dinamiche di radice di pianta è essenziale per migliorare l'efficienza di utilizzo delle risorse dei sistemi agricoli e aumentare la resistenza delle cultivar di raccolto contro stress ambientali. Un protocollo sperimentale è presentato per RGB e l'imaging iperspettrale degli apparati radicali. L'approccio utilizza rizobox dove le piante crescono in terreno naturale sopra un tempo maggiore per osservare completamente sviluppati sistemi della radice. Impostazioni sperimentali sono esemplificate per valutare rhizobox piante sotto stress idrico e studiare il ruolo delle radici. Un'installazione di imaging RGB è descritto per quantificazione economico e veloce lo sviluppo delle radici nel tempo. L'imaging iperspettrale migliora la segmentazione radice dallo sfondo del suolo rispetto alla soglia di colore basata RGB. Il punto di forza dell'imaging iperspettrale è l'acquisizione di chemiometrica informazioni sul sistema suolo-radice per comprensione funzionale. Questo è dimostrato con mapping contenuto di acqua di alta risoluzione. Formazione immagine spettrale, tuttavia, è più complessa in immagine acquisizione, elaborazione ed analisi rispetto all'approccio RGB. Una combinazione di entrambi i metodi in grado di ottimizzare una valutazione completa dell'apparato radicale. Per il contesto della ricerca fisiologica phenotyping e pianta della pianta sono riportati esempi di applicazione l'integrazione di radice e tratti fuori terra. Ulteriore miglioramento dell'imaging di radice può essere ottenuto ottimizzando la qualità di immagine RGB con un'illuminazione migliore utilizzando diverse fonti di luce e di estensione dei metodi di analisi di immagine per dedurre il proprietà di zona di radice da dati spettrali.

Introduzione

Le radici forniscono diverse funzioni essenziali per piante come deposito di assimila, ancoraggio delle piante terrestri nel suolo e l'assorbimento e il trasporto di acqua e sostanze nutritive¹. Da un punto di vista evolutivo, la formazione degli assi di radice è considerata un presupposto fondamentale per l'origine della terra piante². Nonostante questo ruolo importante, storicamente le radici hanno occupato solo una posizione marginale nella ricerca biologica. In tempi più recenti, tuttavia, vi è crescente interesse scientifico nei sistemi di radice di pianta come evidenziato nella Figura 1.

Figura 1: Rilevanza degli studi radice in scienze botaniche.
Numero di radice studi collegati come percentuale di tutti gli studi di pianta pubblicati in riviste SCI negli ultimi decenni. Risultato da Scopus utilizzando parole chiave "impianto" e "pianta e radice" della ricerca. Clicca qui per visualizzare una versione più grande di questa figura.

Due motivi principali possono essere ipotizzati per sottendono i recenti progressi nella ricerca di radice. Prima vegetazione terrestre è esposta a stress ambientali più frequenti a seguito di cambiamenti globali³. Nel contesto della produzione agricola si stima che globalmente circa il 30% della superficie agricola sono limitati dall'acqua e fosforo^4,5. Riduzione dello stress di resa delle colture sono delle ragioni principali per le lacune di rendimento significativo che ammontano globalmente a più basso 50% di produttività potenziale per rainfed agro-ecosistemi⁶. Oltre alla disponibilità di risorse in esaurimento, questo è anche legato all'efficienza di uso di risorsa scarsa, cioè insufficiente capacità di una pianta di sfruttare le risorse disponibili⁷. Questo si traduce in perdite di risorse mobili quali nitrati che possono influenzare negativamente altri ecosistemi. L'efficienza di uso corrente di azoto globale, ad esempio, è stimato a 47%⁸. Uso migliore delle risorse efficienza tramite i metodi di miglioramento della gestione e cultivar è pertanto di grande importanza per entrambi sostenuta crescita dei prodotti agricoli, anche per quanto riguarda la sostenibilità ambientale. In questa pianta contesto radici sono considerate di essere un obiettivo chiave per colture migliorate e ritaglio sistemi^9,10.

Una seconda importante background per il recente interesse nelle radici della pianta è progresso tecnologico nei metodi di misurazione. Metodi di radice lunga sono stati limitati da due sfide chiave: per la misurazione delle radici da piante che crescono nel terreno dovevano essere isolato per quantificazione, principalmente mediante lavaggio¹¹, perturbi la disposizione architettonica degli assi di radice. Osservazione di radice in situ mediante scavo metodi, quindi conservare la naturale posizione delle radici nel terreno, sono stati utilizzati per Descrizione botanica¹². Ancora sono molto che richiede tempo e quindi non soddisfano i requisiti di velocità effettiva del sistema della radice strutturale-funzionale comparativa analisi. D'altra parte metodi ad alta resa per la radice architettura misura principalmente sono state fatte su diete artificiali e per semenzale piante¹³ dove l'estrapolazione all'ambiente naturale crescita delle piante è discutibile¹⁴.

Il recente boom di ricerca principale è collegato strettamente all'anticipo del metodi¹⁵di imaging. Approcci in studi di radice di imaging possono essere approssimativamente raggruppati in tre tipi. Prima ci sono metodi 3D ad alta risoluzione come il CT e MRI¹⁶. Questi metodi sono più adatti per lo studio dei processi di interazione delle radici delle piante con il suolo, come siccità indotta xilema embolia¹⁷. In genere vengono applicati a comparativamente piccoli campioni dove permettono osservazioni dettagliate. Viene fornito un confronto del CT e di MRI per vasi di dimensioni diverse e l'imaging di radice bene in¹⁸. In secondo luogo, ci sono di alto-rendimento imaging metodi^19,20. Questi metodi sono per lo più basata su comuni RGB imaging 2D di radici che crescono su diete artificiali (gel, carta di germinazione) dove il contrasto elevato consente una dissezione comparativamente semplice tra radici e sfondo. Essi sono adatti per il confronto di throughput elevato tra tratti di radice semenzale di genotipi differenti colture sotto crescente artificiale standardizzata circostanze¹³. Tra questi due approcci sono metodi rhizobox: usano imaging 2D di radici che crescono nel terreno nel più lungo periodo di tempo ed hanno velocità effettiva media^21,22. Una sfida risale all'imaging di radice (2D) è quello di catturare anche indicatori di funzionalità principale oltre a descrizione della struttura²³.

Nel presente documento presentiamo i protocolli sperimentali per l'imaging rhizobox cresciuto sistemi della radice mediante (i) una semplice ed economico su misura RGB imaging installazione e (ii) una configurazione più complessa di imaging NIR. Risultati di esempio ottenuti da queste due configurazioni sono mostrati e discussi nel contesto della pianta phenotyping e ricerca fisiologica della pianta.

Protocollo

1. Rhizoboxes for plant growth

NOTE: The experimental system uses rhizoboxes to grow plants for root imaging. First the design of the boxes and the substrate used are described, and then details on the filling procedure are given.

1. Design of rhizoboxes
   1. Create rhizoboxes (Figure 2) with a back plate and side frames made of grey PVC with 15 mm strength. Use a box size of 300 mm x 1000 mm. For the front window, use 6 mm mineral glass which is attached to the PVC frame by metal rails being screwed into the side walls.
   2. Create three holes on the bottom frame to allow drainage of excess water. These holes can be optionally closed by plastic screws.
   3. Before filling, adapt the inner diameter of the rhizobox (between 10 mm and 30 mm) by inserting PC multiwall sheets. An inner space of 10 mm is recommended for most crops to reduce the weight (rhizobox weight without soil is 13.2 kg) of the entire system.

Figure 2: Rhizobox experimental system and its components.
Left figure shows the dimensions of a rhizobox and right figure its single components, being a grey PVC back plate with side frame, a front mineral glass, multiwall sheets for variable inner diameter and metal angles to fix the front glass to the back compartment. Please click here to view a larger version of this figure.

2. Substrate
   1. Fill the rhizoboxes with field soil (for this experiment: silt loam top soil from a calcareous chernozem) sieved to 2 mm particle size.
   2. Open the rhizoboxes for filling the substrate into the inner compartment (back plate with side frames) in horizontal position using pre-wetted substrate. Fill horizontally to avoid layering and segregation between fine and coarse particles which occurs when filling in vertical position by pouring the substrate through the upper opening.
   3. Pre-wet the substrate before filling. Depending on the type of substrate (particularly on its silt and clay content), do not exceed a water content of 0.12-0.18 cm³*cm^-3 to avoid smearing and structure degradation. Add the difference between pre-mixing and target water content after filling the substrate into the rhizoboxes.
      NOTE: Filling with (oven-)dry substrate and subsequent addition of the entire water is not recommended as is can result in strong settlement of the substrate and formation of large cracks.
3. Step by step filling example
   1. Define the target water content. Here it is initially set at 80% plant available water (PAW) where plants do not suffer from any water shortage.
   2. Determine field capacity (FC) and permanent wilting point (PWP) of the substrate. Here obtain FC using a PVC tube of equivalent height (100 cm) as the rhizoboxes.
      1. Close the tube at the bottom with a stubble having small drainage holes, add 1 cm of gravel to avoid closure of the holes from finer substrate and fill the substrate to the same bulk density as used for the rhizoboxes (1.3 g cm^-3).
      2. Saturate the tube with water until drainage occurs and leave it for two days to equilibrate (the resulting water content is per definition equivalent to field capacity), while covering the upper opening with cling-film to avoid evaporation. The field capacity value achieved for the soil in this experiment was 0.357 cm³ cm^-3.
         NOTE: Water content at PWP has to be known in advance from standard soil physical methods (e.g. pressure plate measurements²⁴) or from texture based pedotranfer functions²⁵. Here it is equal 0.12 cm³ cm^-3 for the soil used.
      3. When measuring FC by pressure plate extraction too, take the water content at a matrix potential of h=-100 hPa and not h=-330 hPa in order to correspond to the rhizobox geometry (height of 100 cm = 100 hPa).
      4. Calculate the water content (WC) at 80% PAW: WC (cm³ cm^-3) = 0.80 (FC-PWP) + PWP. For the hydraulic limits of the soil used here this gives a volumetric water content at 80% PAW of 0.31 cm³ cm^-3.
      5. Calculate the water amount for a rhizobox volume of 2850 cm³ (30 cm width, 1 cm inner space, 95 cm in height, with top 5 cm keeping free of substrate for watering). This gives a water volume of 883.5 cm³ equal 883.5 g for the density of water being 1.0 g cm^-3 at 20 °C.
   3. Define the bulk density (d_b) to fill the rhizoboxes. Here, set at 1.3 g cm^-3 corresponding to values typically found in agricultural field soils. The amount of dry substrate required to fill a rhizobox volume of 2850 cm³ at this d_b equals 3705 g of dry soil.
   4. Pre-wet the dry soil to a gravimetric water content of 0.108 g g^-1 (equal to a volumetric water content of 0.14 cm³ cm^-3) by adding 400 g of water for 3705 g of dry soil and mix it gently to obtain a homogeneous water distribution. Manually disrupt larger aggregates to keep the particle size ≤ 2 mm.
   5. Fill the pre-wetted soil into the opened rhizoboxes and compact it gently using a polystyrene sheet (30 x 10 x 1.5 cm) to cover the inner volume of the box, thereby resulting in a homogeneous d_b of 1.3 g cm^-3.
   6. Add the remaining amount of water (483.2 g) to achieve the target water content of 0.31 cm³ cm^-3 by spraying onto the surface with a spray bottle. Ensure small drop size to avoid surface structure degradation and homogeneous wetting. Keep the box on a balance during spraying to monitor the amount of water actually added to substrate.
   7. Let the water redistribute for 10 minutes and then press the glass onto the surface and fix it with the side metal rails. The average final weight of rhizoboxes with wetted substrate was 17818 ± 68 g (13230 g rhizobox weight + 3705 g dry soil + 883 g water).
      NOTE: The homogeneous water content filled in the horizontal boxes will redistribute when the boxes are set in their final position according to the resulting potential gradient. This is a physical process in all plant growth pots according to their geometry (height) and experimenters should be conscious on their pot hydraulics²⁶.

2. Climate room setup

1. Equip the climate room (Figure 3) with 8 LED lamps providing homogeneous illumination of 450 μmol m^-2 s^-1 with spectral peaks at 440 (blue) and 660 (red) nm for optimum plant growth.

Figure 3: Climate chamber with rhizoboxes for stress experiment.
(A) Left view of entire chamber with LED illumination, weather station and PC (here for logging leaf hygrometers); right view with a close-up of the metal frame holding rhizoboxes in 45 ° inclination and wooden plates used to shied rhizobox glass window against light. (B) Stress experiment with sugar beet combining four stages with stress due to different atmospheric demand (high/low) and soil water availability (high/low). Green bars of mean stomata conductance give an indication of plant stress response. Please click here to view a larger version of this figure.

2. Set the ambient parameters according to plant/experimental needs. Here, use 14 hours light and 10 hours dark for illumination. During plant establishment and before stress treatments started, set the temperature to 20° C during day and 15° C during night and keep relative humidity at 50 ± 8%.
3. Put the rhizoboxes at an inclination of 45° using an adequate metal framework. This maximizes root growth towards the glass surface due to gravitropism.
4. Cover the glass window by a wooden plate to keep the root zone in dark and avoid algae growth due to light penetrating through the glass surface.

3. Sugar beet example setup and treatments

1. Pre-germinate sugar beet seeds on wet filter paper for three days at 20 °C in an incubator until the radicle emerges. This ensures seeding with viable plants.
   NOTE: Pre-germination is not required for plants with high germination vigor, thereby avoiding the risk of damaging the radicle at seeding. For sugar beet seeds with a thick pericarp, however the risk of non-viable seeds is high and pre-germination significantly accelerates the emergence of the radicle compared to direct seeding into soil.
2. Drill a small hole to about 1.5 cm soil depth in the middle of the rhizobox with a screwdriver, position one seed in it using tweezers with the radicle oriented downwards and next to the glass window (this improves the initial visibility) and gently cover it with soil.
3. Add a 0.5 cm layer of fine gravel (2-4 mm) on top of soil to protect the soil aggregates from slaking during irrigation and reduce evaporation losses. To facilitate emergence, keep the soil surface free of gravel where the seed has been positioned.
4. Add 10 g of water to enhance establishment.
5. During establishment and early growth until experimental stress treatments start, irrigate the rhizoboxes every 2-4 days to keep the initial moisture content of 80% PAW.
   1. Determine the amount of irrigation water required by weighing the rhizoboxes and adding water until achieving the initial weight of each individual box. For manual irrigation use a pipette to avoid surface structure degradation during watering.
6. Arrange the rhizoboxes according to the established design in the climate room. The protocol reported here is based on an experiment with six sugar beet cultivars in five replicates in a completely randomized design (CRD).
   1. Reposition rhizoboxes each time when taking them out of the metal holder for weighing and watering. This avoids any effects of residual (light) inhomogeneity inside the climate room.
7. Define the time for onset of stress treatments and the type of stress. The following settings (Figure 4) are used here.
   1. Start measurements at BBCH 15 (five leaves unfolded) with roots covering around 75% of the rhizobox depth and the canopy being sufficiently developed for measurements at the leaves. Keep each stage for at least three days to ensure adaptation to the new settings and make measurement.
   2. For the non-stress observations keep the initial settings with optimum soil moisture (80% PAW) and ambient conditions (20° C/15° C temperature, 50 ± 8% rH) giving a daytime vapor pressure deficit (VPD) of 1.28 ± 0.1 kPa (cf. Figure 3 C).
   3. Raise the atmospheric demand to a daytime VPD of 2.45 ± 0.4 kPa by increasing the temperature to 27°C/20°C and decreasing rH to 35%, while keeping soil moisture at 80% PAW.
   4. Subsequently dry down the rhizoboxes to 40% PAW equivalent to a water content of 0.215 cm³ cm^-3 by withholding irrigation. Reset the initial ambient conditions with low atmospheric demand (VPD of 1.28 kPa).
   5. Combine stresses increasing the atmospheric demand to a VPD of 2.45 kPa and keep soil moisture at 40% PAW.

4. Root imaging methods

1. Combine imaging methods to make use of their respective advantages and depending on the information targeted.
   1. Apply RGB imaging in the VIS range to track root growth, architecture and morphology over time which implicitly requires frequent measurement. Advantages of RGB imaging are (i) low costs, (ii) rapid image acquisition, (iii) low requirements of hard-disk space (image size: 48 MB) and (iv) high resolution (3648 x 5472 pixels).
   2. Use hyperspectral imaging (HSI) in the NIR range when chemometric features of root and soil are required. Advantages are (i) spectral features for segmentation between roots and soil background and (ii) access to physico-chemical system properties (e.g. soil water content, root age). Disadvantages are (i) higher scanning time (about 16 minutes per rhizobox), (ii) large size of datasets (13.7 GB per rhizobox image) and (iii) lower resolution of the NIR camera (320 x 256 Pixel), and (iv) higher complexity of data analysis.
2. RGB root imaging
   1. Use an imaging box (Figure 4) that shields from ambient light and fixes the camera position consisting of a metal frame with a width of 1 m and a height of 1 m with side walls lined with pressboards. At the front end, fix the camera at two positions at a distance of 80 cm from the rhizobox.
      1. Attach a tapeline on the rhizobox frame with transparent adhesive tape and position the rhizobox in the holder of the imaging box.
      2. Illuminate the rhizobox using four 24 W fluorescent light tubes attached at a distance of 80 cm from the rhizobox. Also, mount four 15 W UV tubes at 20 cm from the rhizobox as alternative illumination making use of root auto-fluorescence in case of low contrast between root and (bright colored) substrate background.
      3. Take two images (top and bottom position) to cover the upper and the lower half of a rhizobox with an overlap of about 3 cm.
   2. Acquire RGB images with a digital single-lens reflex camera which is fixed by quick release plates on the respective positions of the imaging box.
      1. Apply the following settings when using the fluorescent light tubes. Adapt these example settings for any changes in dimensions and illumination as well as the camera model.
         1. Turn off autofocus and stabilizer on the camera objective. Set camera to manual mode.
         2. Set ISO speed to 500. Set shutter speed to 13. Set Aperture to 5.6.
         3. Turn off the mirror lock. Set the white balance to Auto White Balance.
      2. Use the following settings for illumination with UV tubes:
         1. Turn off autofocus and stabilizer on the camera objective. Set camera in manual mode.
         2. Set ISO speed to 1000. Set shutter speed to 13. Set Aperture to 5.6.
         3. Turn off the mirror lock. Set the white balance to Fluorescence.
   3. Merge the RGB images from top and bottom of the rhizobox into a single image a photo editor (e.g., Adobe Photoshop). Use the tapeline at the side frame of the rhizoboxes and control overlapping objects (root and soil features).
      1. Copy the two separate images, each with pixels size of 3648 x 5472, into a new file of size 3648 x 10944 pixels and white background.
      2. Reduce the layer opacity for one image to 60% and align overlapping parts of the images (tapeline, objects). Thereafter restore layer opacity to 100%.
      3. Based on the ruler on the image add two red lines of exactly 1 cm to the top of the image where no roots are present. These lines are later used at image analysis to scale the image to correct length dimensions.
      4. Merge all layers and remove parts of the image outside the soil filled window with the cropping tool.
      5. Save the image as tiff-file for further analysis.
      6. In case of images with UV illumination, reduce colors to greyscale before saving using the toolbar Adjustments-Black & White, and selecting the predefined High Contrast Blue filter.
   4. For segmentation between roots and soil background as well as for quantification of root traits of interest (e.g. length, surface, diameter, branching) use any root analysis software (see www.plant-image-analysis.org for available tools). Here WinRhizo is used.
      1. Open the rhizobox tiff-image the software.
      2. Calibrate the length scale of the image using the scaling bars added to the image.
      3. Select Based on Color in the Analysis menu where selecting Root & Background Distinction.
      4. Define a calibration file with color classes corresponding to roots and soil (background). For the example images used here (cf. Figure 8) three root color classes (old laterals, young laterals, tap root) and three soil color classes are defined.
      5. For the greyscale images (UV-illumination) select (i) Based on grey levels, and (ii) Pale Root on Black Background in the Root & Background Distinction menu and use a local Lagarde intensity threshold for segmentation.
      6. Open a data file where results are saved.
      7. Run the analysis and subsequently control whether there are regions (e.g. at the edges) which are mismatched. In this case define an exclusion region and restart the analysis. For roots not classified, add additional color classes and restart the analysis. For elements wrongly classified as roots, activate/increase the Debris & Rough Edges filtering options.

Figure 4: Imaging box to acquire RGB rhizoboxes pictures.
Left view of front side where rhizoboxes are attached for imaging with light sources inside; right view of backside where the camera is mounted. Please click here to view a larger version of this figure.

3. Hyperspectral root imaging
   1. Hardware setup
      1. Use a hyperspectral root imaging system (Figure 5) consisting of (i) a thermo-electrically cooled 14-bit monochrome NIR camera with a spectral range from 900 nm to 1700 nm, 320 by 256 pixels and a frame rate of 100Hz and (ii) an imaging spectrograph with a spectral range 900 nm to 2500 nm and a spectral resolution of 3.6 nm. Arrange a halogen line illumination source (four 50 W halogen spots) in a 45°/-45° geometry. Mount the imaging sensor on a two-axis positioning system. The scan window has a size of 240 x 1000 mm, i.e. 30 mm at each edge of the rhizobox are not covered by the image.
      2. Control the system by a Matlab script for (i) white and dark standard acquisition, (ii) setting of camera integration time, (iii) selecting spatial (pixel size 0.1 mm; pixel size 1.0 mm) and spectral resolution (all 222 spectral bands with a resolution of 3.6 nm; smoothed spectrum with 54 bands and a resolution of 14.8 nm), and (iv) defining the scan region on the rhizobox.
      3. Save images as SIF-files. To avoid problems during saving of large files, subdivide each scanned image stride (9 strides per rhizobox) into four segments (three of 300 mm length, one of 100 mm length) and save separately with a unique file name consisting of stride number (1 to 9) and part (1 to 4) as well as date and time (YYYY.MM.DD HH:MM:SS). An entire rhizobox scan requires a hard-disk space of 13.7 GB.
   2. Image acquisition and analysis.
      NOTE: Figure 6 shows the steps of image acquisition, segmentation and analysis.
      1. Image acquisition comprises selection of camera setting for optimum image quality and definition of scan parameters.
         1. Determine the camera integration times for the rhizobox scan and the white standard in the camera software.
            1. Open the imaging GUI and move the camera to a position of the rhizobox where roots are present.
            2. Adjust the integration time of the camera targeting a light object (i.e. root) in a way that approximately 85% of the full dynamic range of the camera is used on the histogram displayed by the software. Repeat for the white standard by moving the camera positioning system to target the white standard. Then close the camera software.
         2. Open the Matlab Imaging GUI and make all settings for the current rhizobox scan. For the data reported here, use the following settings:
            Integration time white standard: 1000
            Integration time rhizobox: 4000
            Spectral resolution: Full resolution (i.e. 222 narrow-range spectral bands)
            Full spatial resolution (pixel size of 0.1 mm)
            1. Acquire the dark and white standards before each imaging run, e.g. once a day. The dark standard represents the camera noise, while the white standard gives the maximum reflectivity. These data are required for image normalization during pre-processing.
            2. Define whether the entire rhizobox or only part of it is scanned. For the present case entire rhizoboxes are imaged. Then start the scan.
      2. Process the image with a Matlab script. Operations performed by the script are described.
         NOTE: Scripts are currently in an undocumented version and can be obtained from the corresponding author. After proper documentation, they will be available for download from the website of the corresponding author's institution (www.dnw.boku.ac.at/pb/).
         1. Compose an entire image stride from the rhizobox center (containing roots) merging the four parts of the stride.
            NOTE: At this stage it is neither necessary nor recommended to use a spectral image of an entire rhizobox (i.e. all 9 strides) as the file size will make each calculation step in Matlab very time consuming and the information contained in one central stride is sufficient for the first steps of image analysis.
         2. Normalize the image using the acquired dark and white standards and taking into account the different integration times of white standard and rhizobox scans which are saved automatically during scanning in a file.
         3. Optionally apply a smoothing filter to remove noise from the image. The script currently offers 3x3 kernel median filtering and multiple scatter correction. For the image evaluation presented here, no filters are applied.
         4. Display the image at all recorded spectral bands to obtain a first insight and decide a wavelength to be displaying for selecting regions of interest (cf. image segmentation).
      3. Perform segmentation between roots and soil background in a separate script with the following steps.
         1. Select regions of interest (ROI) for root and soil to find spectral features for segmentation. Use the freehand selection tool to mark a ROI on the image displayed at a wavelength previously identified with good contrast between roots and soil. Here, use three ROIs on the root (old and young laterals, tap root) and two ROIs in the soil (dry, wet region).
         2. Display a rectangle with the selected ROIs and the remaining part as a black mask and visualize the selected ROI image at all wavelengths.
         3. Remove all lines in the image matrix containing pixels of spectral intensity = 0 (black pixels).
         4. Fuse the root and soil ROIs into one foreground (root) and one background (soil) matrix for segmentation.
         5. Search spectral bands (intensity of single spectra or spectral ratios) providing the best separation between root foreground and soil background²⁷. Quantify the distinction between the resulting pixel histograms for root and soil using Bhattacharyya distance²⁸.
         6. Select a threshold intensity value separating the histograms.
         7. Create a binary image by applying the selected threshold to the original image. This sets all pixels having smaller intensity than the threshold to zero and those with higher intensity to one (done automatically be the script).
         8. Save the binary image as tiff-file.
      4. Open the binary image and analyze root traits. Select (i) Based on grey levels, and (ii) Pale Root on Black Background in the Root & Background Distinction menu and use global intensity threshold (the image is already binarized).
      5. For mapping the water content from the hyperspectral data acquire a calibration dataset and apply a calibration equation to a rhizobox image.
         1. Subdivide a rhizobox into 5 cm compartments using polystyrene sheets to fill them with soil (same substrate and d_b as used for the experiment) at different water contents (Figure 8).
         2. Calculate the respective amounts of water to be mixed with soil and fill the compartments (same procedure as described in 1.3. for the entire rhizobox).
         3. Scan the calibration rhizobox with the same settings as used for the planted rhizoboxes.
         4. Perform the following steps using a script.
            1. Merge the four parts of a stride from the water calibration box to one stride and normalize it with dark and white standards.
            2. Select rectangular boxes at each compartment with different water content and save them in a structure array.
            3. Determine the spectral feature that best separates the water content compartments. This is done with a search algorithm for the global maximum of the intensity differences between mean spectra of adjacent water contents.
            4. Calculate the mean spectral intensity value for this feature for each water content compartment of the calibration rhizobox.
            5. Fit a regression equation through the directly measured water content and the respective spectral quantifier.
            6. Apply the regression equation to each pixel not classified as root on a rhizobox image and for the spectral feature (band) determined above which best relates to water content.

Figure 5: Hyperspectral root scanner.
The main components of the scanner are indicated. The small picture shows the camera during imaging of a rhizobox. Please click here to view a larger version of this figure.

Figure 6: Steps in hyperspectral root imaging.
Hyperspectral root imaging consists of three mains steps being (i) image acquisition, (ii) image segmentation and (iii) analysis of the spectral data. Please click here to view a larger version of this figure.

Figure 7: Rhizobox for water calibration.
The rhizobox contains compartments with substrate at different water content which are subdivided by polystyrene sheets. Germination paper at the dry compartments ensures that soil particles do not rinse into neighboring compartments. Please click here to view a larger version of this figure.

5. Application examples

NOTE: Quantitative root information is applied in the context of plant phenotyping (cultivar comparison) and for plant physiological research. The following aboveground data are reported to exemplify these cases.

1. Leaf area: Measure leaf area non-destructively at selected stages during the experiment via the length and width of leaves as a proxy. Alternatively canopy images can be used²⁹.
   1. At the end of the experiment measure leaf length and width together with area of clipped leaves using a leaf area meter. Calibrate the non-destructive method applying a regression equation to the data pairs.
2. Dry matter: At the end of the experiment, measure aboveground dry matter by clipping the plants with scissors and dry them for 24 hours at 105 °C in an oven.
3. Stomata conductance: Measure stomata conductance with a leaf porometer. Before measurement, keep the device for at least one hour in the climate chamber to allow sensors equilibrate with ambient conditions and calibrated the device each time when ambient settings in the climate chamber are changed. Take measurements from at least three leaves per plant.

Risultati

Example results are presented for root segmentation based on RGB and HS imaging. For spectral imaging an example of high resolution water mapping is provided. Finally results are shown that demonstrate the scientific context where image based root data are applied.

RGB based root measurement

Figure 8 shows an RGB root image time series of sugar beet cultivar Ferrara. The images reveal some artefacts from inhomogeneous illumination of the rhizoboxes, with brighter areas along the left side and different brightness at the overlapping area between top and bottom images.

Figure 8: Root growth time series from the RGB imaging.
Pictures show the sugar beet cultivar Ferrara at different days after sowing (DAS). The images show some artefacts due to non-homogeneous illumination at the left side of the image and between top and bottom images. Scale bars, 2 cm. Please click here to view a larger version of this figure.

Figure 9 provides details on root segmentation based on color thresholding for cultivar Ferrara at day after sowing (DAS) 35. As a reference (Figure 9A), a binary image is used where all roots were manually tracked with a Graphic Tablet. The time required for manual tracking of the entire, fully developed, dense sugar beet root system was around four hours. Figure 9B gives a detailed view on a selected area at the top of the image where old lateral roots are present. Here several root axes are not classified by the color threshold. At the bottom (Figure 9C) on the contrary, where white young roots are predominant, the color based segmentation properly classifies all root axes. The binarized root system (Figure 9D) shows a black area at the left side from the illumination artefact which was defined as exclusion region before running quantitative analysis. Figure 9E shows the corresponding pixel histograms of selected features (roots vs. soil) for the red channel of the RGB image from Ferrara at DAS 35. The root pixels (blue color) clearly show three peaks corresponding to bright young laterals, dark old laterals and tap root. The overlap between the old laterals and the soil background is very strong, leading to unclassified root axes (cf. Figure 9B).

Figure 9: Root segmentation using a color threshold.
(A) Manually segmented root system using a Graphic Tablet, (B) area with poorly segmented old root axes in the top and (C) properly segmented young axes in the bottom of the image, and (D) binary image obtained from color based thresholding. (E) Pixel histograms for selected features of the RGB image. Roots are represented by the blue bars with different root types indicated; soil is represented by the red bars. Scale bars in A and D, 2 cm; scale bars in B and C, 1 cm. Please click here to view a larger version of this figure.

The resulting total visible root length quantified for the manually segmented reference image is 1534.1 cm, while the automatized, color based segmentation gives a total root length of 1427.6 cm.

Greyscale images from UV-illumination do not provide an advantage in the case shown here and performed worse compared to color thresholding (root length: 1679.7 cm). Old roots could not be segmented, and there was more noise in the image, probably due to lower light intensity of the UV lamps. However, in case of young roots with high auto-fluorescence and a bright background substrate, UV-illumination can still be an option as shown by an image obtained from another experiment where sand was used as background substrate (Figure 10).

Figure 10: UV illumination to visualize roots on bright background.
Example from a durum wheat root system growing in a rhizobox filled with quartz sand. The rhizobox is imaged with illumination for (A) UV light and (B) fluorescent (day) light. Scale bars, 2 cm. Please click here to view a larger version of this figure.

HSI based root measurements

Figure 11 provides the mean spectra for three root ROIs (old and young lateral, tap root) and two soil ROIs (top and bottom of rhizobox).

Figure 11: Mean spectra of root and soil.
Spectra from regions of interest (ROIs) on the root (three root types) and in the soil (top and bottom of the rhizobox). The ROIs are selected to determine an optimum segmentation criterion between root and soil. Please click here to view a larger version of this figure.

It is evident that the tap and young lateral roots differ substantially from the background in intensity of most spectral bands. For the old laterals the intensity differences are much lower. A feature that can be inferred visually is the different slope of the spectrum around water absorption region (1450 nm). Here the slope of root spectra is higher compared to soil spectra. Furthermore a change of tap and young lateral spectra in the region around 1100 nm can be identified that does not occur in the old laterals.

Figure 12A shows the result from the search algorithm identifying a spectral ratio with strongest foreground-background contrast. The ratio of spectra at 1476 nm to 1076 nm provides the best separation between roots and soil. The resulting histogram of root foreground and soil background pixels is shown in Figure 12B. Although there is some overlap, most pixels are clearly separated from the soil background. Fitting a bimodal Gaussian curve through the histogram and using Bhattacharyya distance for quantification, a value of 7.80 is obtained. A value higher 3.0 indicates strong image contrast allowing reliable separation²⁸.

Figure 12: Difference in reflectance between root foreground and soil background for different spectral band ratios and pixel histogram at spectral ratio used for segmentation.
(A) Bright colors (yellow) show high contrast between foreground and background, dark colors (blue) show low contrast. The first 15 bands have been removed because of noise. The red lines indicate the band ratio with highest contrast. (B) Pixel histogram of roots (blue) and soil (red) at segmentation spectral ratio. Blue bars represent the root and red bars the soil. The intensity value corresponds to the ratio of spectral band 160 to spectral band 49. Please click here to view a larger version of this figure.

The binary image (Figure 13) is created by applying a global intensity threshold of the identified spectral ratio at a value of 1.008 calculated from the histogram distance²⁷. Analysis of root length of this image gives a total length of 1557.3 cm which represents an error of only 1.5% compared to the manually tracked reference image.

Figure 13: Binary image of the root system of sugar beet cultivar Ferrara.
The image is obtained by applying a global spectral threshold. Scale bar (bottom left corner), 2 cm. Please click here to view a larger version of this figure.

Although root segmentation has improved using spectral information compared to color based information, the main intention of HS imaging is analysis of chemometric image properties. This is exemplified via mapping the water content of a rhizobox image.

Figure 14A shows the mean spectra of the compartments in the calibration rhizobox (cf. Figure 7) filled with soil of different water content. The shape of spectra is similar between the compartments, i.e. here a spectral ratio does not necessarily provide a more stable classification criterion. Thus intensity at a single spectral band (1680 nm), where the average difference between adjacent water contents is maximized, is identified as best separating criterion. The resulting pixel histograms for this spectral wavelength are shown in Figure 14B.

Figure 14: Spectral features for water content calibration.
(A) Mean spectra of nine water compartments from the calibration rhizobox with different water contents; (B) Pixel histograms for the water compartments at band 216 where average distance between neighboring compartment is maximum. Please click here to view a larger version of this figure.

The relation of the average pixel intensity at 1680 nm and the measured water content is shown in Figure 15.

Figure 15: Relation of spectral reflectance and volumetric water content.
The figure shows data pairs of measured water content and spectral reflectance with empirical curves (linear and exponential) fit to the data excluding the highest water contents (red triangles). Please click here to view a larger version of this figure.

Differentiation of higher water contents from spectral intensity becomes difficult. A significant regression (either linear or exponential) with high R² can be fit to water contents up to around 0.30 cm³ cm^-3. Wetter soil conditions cannot be reliably predicted by the intensity value. Similar behavior of an exponential relation between reflectivity and water content with a decreasing response to water contents higher 0.30 cm³ cm^-3 was also found in other studies³⁰.

A rhizobox image with fine mapping of water content is shown in Figure 16. Four aspects have to be remarked. First, a region of lower water content can be seen in the rooted parts of the rhizobox. Second, strongest depletion is concentrated in the vicinity to single root axes. Third, depletion zones also occur where no root axes are visible on the surface, indicating regions where roots are hidden in soil. Fourth, water mapping without further image-processing results in a patchy appearance due to the aggregated soil. This can indicate inhomogeneous water content distribution at the aggregate scale, but also surface morphology effect on image quality. Chemometric image-processing techniques are an option to overcome such morphological effects in spectral images³¹, but are not implemented so far in the Matlab scripts used here.

Figure 16: Water content mapping on a rhizobox.
The dark blue colours represent regions of high water content, green to red areas show regions with low water content. The plant root is overlaid on the image in black. Scale bar (bottom left corner), 2 cm. Please click here to view a larger version of this figure.

Application examples

Figure 17 relates quantitative root traits from image analysis with aboveground measurements.

Figure 17: Typical application examples for root data.
(A) and (B) show root information used for aboveground-belowground plant characterization in a phenotyping context. (A) represents root growth from the sugar beet cultivar Ferrara, (B) compares six rhizobox grown sugar beet cultivars using leaf-to-root area ratio (data from one replicate). (C) and (D) are functional relations between traits as found in plant physiological research. (C) shows the influence of leaf-to-root area ratio on dry matter production and (D) the relation of root surface area to stomata conductance. Please click here to view a larger version of this figure.

Figure 17A and 17B are relevant for phenotyping focusing on comprehensive aboveground and belowground plant characterization. Figure 17A shows root growth of sugar beet cultivar Ferrara (cf. Figure 8 for images). Expansion of the root system indicates the capacity of a cultivar to explore the soil volume in a given time span of the vegetation period. Figure 17B shows leaf-to-root surface area ratio of six sugar beet cultivars, providing a descriptor for the balance between plant supply (root) and demand (leaf).

Figures 17C and 17D give examples for functional relations of interest in physiological research. In Figure 17C leaf-to-root surface area ratio is related to dry matter formed during the experiment, indicating the predominant role of the assimilating surface as a limiting factor for dry matter accumulation. The lack of significance in spite of a comparatively high R² is related to the low number of paired data (n=6) used here. Figure 17D reveals that cultivars with higher root surface area (improved uptake) have an average higher stomata conductance over the course of the experiment. The higher root area apparently sustains water extraction, thereby prolonging stomata opening.

Discussione

I protocolli prevedono due approcci complementari terreno coltivato imaging system di root. Un passaggio fondamentale per ottenere risultati sperimentali affidabili si sta riempiendo di rizobox che deve garantire un livello di substrato uniforme ed omogeneo nel vetro anteriore per offrire suolo-radice stretto contatto nella finestra di osservazione ed evitare vuoti d'aria. Questo è il motivo principale per utilizzare terra comparativamente fine setacciata di < 2 mm: più grandi aggregati provocare la morfologia superficiale superiore alla finestra di osservazione con i vuoti tra gli aggregati. Oltre ad un elevato rischio di disidratazione di punta di radice, questo richiede anche più complesse tecniche di elaborazione di immagini per acqua mappatura³¹.

Modifiche del protocollo, pertanto concentrano sul migliore e rapido riempimento di rizobox. Attualmente il tempo di riempimento è circa 30 minuti per scatola. Inoltre l'uso di rizobox con due finestre di vetro per l'imaging da entrambi i lati e modifiche per ottimizzare l'omogeneità di illuminazione per ottenere immagini migliori RGB vengono testati. Ulteriore estensione hardware anche considerare l'integrazione di planare optodi³² , nonché capacità di imaging³³ nel sistema rhizobox. Si tratta tuttavia oltre le attività di aggiornamento corrente.

Modifiche del software di concentrano sulla registrazione automatica dell'immagine per fondere la parte superiore e inferiore RBG immagini³⁴. Hyperspectral imaging avanzate funzione senza supervisione di estrazione si avvicina per²⁸ , nonché metodi di rilevazione più sensibile obiettivo sorvegliato come SVMs³⁵ sono testati. Quindi i dati iperspettrali potenzialmente consentono per la valutazione di più terreno, rizosfera e radice di proprietà³⁶. Inoltre si intende sviluppare un (semi) automatizzato software per rhizobox radice immagini basato su una versione modificata di radice sistema analizzatore³⁷ quantificare morfologiche (lunghezza, diametro, superficie) e tratti architettonici (frequenza ramificazione, ramificazione angoli).

La principale limitazione del protocollo rispetto ad approcci di imaging 3D è la restrizione per la superficie visibile radice e la proprietà di rizosfera. Tuttavia è stato dimostrato che i tratti visibili radice sono un proxy affidabile per l' intero sistema della radice²¹. La tecnica di rhizobox è facilmente combinato con campionamento distruttivo tradizionale (lavatrice) alla fine di una crescita dinamica imaging al fine di convalidare la relazione di visibile vs totale radice sistema tratti. Come questa relazione potrebbe variare tra specie²¹, campionamento distruttivo è consigliabile per garantire affidabile inferenza dai tratti visibili per qualsiasi nuova serie fenotipizzazione con una specie di colture diverse.

Il vantaggio fondamentale del protocollo presentato qui è la combinazione di condizioni di crescita realistiche (suolo), potenziale relativamente alto throughput per la temporaneamente risolto dell'imaging RGB e inferenza sulla funzionalità di radice (ad esempio l'assorbimento di acqua) tramite i dati di radice e rizosfera chemiometrica da imaging iperspettrale. Quindi i metodi supera le restrizioni di inferenza in semenzale elevato throughput e fuori suolo radice imaging metodi¹⁴, mentre permette parzialmente intuizioni profonde che phenotyping processi funzionali con complessità meno sperimentale e un throughput più elevato rispetto ai metodi avanzati 3D¹⁵.

Nei prossimi esperimenti utilizzerà il protocollo per studiare l'effetto di Micorriza sullo sviluppo del sistema della radice e funzionalità dei legumi anche per quanto riguarda le caratteristiche di radice fenotipizzazione di coltura di copertura specie in relazione alla struttura del suolo, di azoto e di carbonio in bicicletta.

Materiali

Name	Company	Catalog Number	Comments
Rhizobox	Technisches Büro für Bodenkultur	Experimental builder
LED Lamps ATUM HORTI 600	Klutronic	AHI10600F
Fluorescent light tube HiLite T5 Day	Juwel Aquarium	86324
UV light tube Eurolite 45cm slim 15 W	Conrad	593384 - 62
Canon EOS 6D	Canon Austria GmbH	8035B024
Adobe Photoshop CS5 Extended Version 12.0 x 32	Adobe Systems Software Ireland Ltd.
WinRhizo Pro v. 2013	Regent Instruments Inc.
Xeva-1.7-320 SWIR camera	Xenics	XEN-000105
Spectrograph Imspector N25E	Specim
Hyperspectral imaging scanner	Carinthian Tech Research AG	Experimental builder	Design and assemblage of Hyperspectral Imaging Scanner and software
Matlab R2106a	Mathworks		Including Toolboxes for Image Processing, Signal Processing and Statistics and Machine Learning
AP4 Poromoeter	Delta-T-Devices
LI-3100C Area Meter	LI-COR
BASF Styradur polystyrene sheets	Obi Baumarkt	9706318	Different types of polystyrene sheets or other material separating differently moistured soil can be used