An evaluation of inexpensive methods for root image acquisition when using rhizotrons
© The Author(s) 2017
Received: 12 August 2016
Accepted: 22 February 2017
Published: 7 March 2017
Belowground processes play an essential role in ecosystem nutrient cycling and the global carbon budget cycle. Quantifying fine root growth is crucial to the understanding of ecosystem structure and function and in predicting how ecosystems respond to climate variability. A better understanding of root system growth is necessary, but choosing the best method of observation is complex, especially in the natural soil environment. Here, we compare five methods of root image acquisition using inexpensive technology that is currently available on the market: flatbed scanner, handheld scanner, manual tracing, a smartphone application scanner and a time-lapse camera. Using the five methods, root elongation rate (RER) was measured for three months, on roots of hybrid walnut (Juglans nigra × Juglans regia L.) in rhizotrons installed in agroforests.
When all methods were compared together, there were no significant differences in relative cumulative root length. However, the time-lapse camera and the manual tracing method significantly overestimated the relative mean diameter of roots compared to the three scanning methods. The smartphone scanning application was found to perform best overall when considering image quality and ease of use in the field. The automatic time-lapse camera was useful for measuring RER over several months without any human intervention.
Our results show that inexpensive scanning and automated methods provide correct measurements of root elongation and length (but not diameter when using the time-lapse camera). These methods are capable of detecting fine roots to a diameter of 0.1 mm and can therefore be selected by the user depending on the data required.
KeywordsFine root elongation rate Flatbed scanner Handheld scanner Smartphone Time-lapse camera
Fine root growth, defined as apical elongation over time [1–3], plays an essential role in the cycling and allocation of carbon and nutrients in ecosystems . Due to the inaccessibility of root systems, special techniques are required to investigate the distribution and dynamics of roots, as well as to estimate belowground carbon budgets [5, 6]. Today, a number of methods have been used to estimate root growth. These methods can be grouped into indirect and direct techniques [5, 7], both of which have advantages and drawbacks. Indirect methods include the use of empirical models , estimations of nitrogen budget and carbon budget . Direct methods can be classified into (i) destructive techniques such as soil coring , sequential soil coring , in-growth cores [5, 10], monoliths [11–13] and soil pits [14–16], and (ii) nondestructive in situ methods including isotope quantification , ‘root windows’ or rhizotrons [18–21] and minirhizotrons [3, 22, 23]. Although there are several criticisms concerning these techniques , rhizotrons and minirhizotrons are considered as efficient approaches and are commonly used to characterize fine root growth [24–26]. Rhizotrons can be used to monitor (from initiation to mortality) specific root segments at frequent time intervals without significantly impacting root processes . However, the drawbacks of these techniques are related to the cost of installation and potential changes in soil hydrology and physics, which would affect estimates of root production [7, 27].
Although many studies on root growth using minirhizotrons have been performed [23, 28], only a small part of the root system can be observed. Techniques for observing root growth include recording root images with digital cameras [23, 29] and rotating scanners (CID, Inc., WA, USA) , but equipment is expensive. Results from contrasting methods on one single species can also be highly variable . The disparity in results obtained from different methods [15, 16, 30–33] has also been attributed to differences in the software used for image analysis [34–36].
The quality of images obtained from minirhizotrons and rhizotrons is extremely important for an accurate quantification of root growth through image analysis. The advantage of rhizotrons over minirhizotrons, is that a variety of inexpensive techniques exist for quantifying root growth in the field. Root systems can be measured by tracing onto a transparent plastic sheet , scanning with a flatbed scanner [30, 32, 34], or a handheld scanner . Scanners have often been considered as the most useful tool for obtaining high quality images, but necessitate the use of a power supply in the field and are not yet fully automated. A detailed comparison of the different types of scanners available has also not yet been performed, especially with regard to the scanners now available as digital applications on smartphones and tablets. Such a comparative study would be highly useful, especially when choosing a particular scanner for a given application and considering its cost, robustness, automation and the quality of the images produced.
With regard to recording automatically images in the field, systems that are independent of an electrical power supply are not yet available, although automated flatbed scanners using 12v batteries for several days have been tested successfully in a teak (Tectona grandis ) plantation in Lao PDR (Maeght, unpublished data). A fully automated method for measuring root elongation in the field would permit studies of growth in poorly accessible areas or with a poor power supply, as well as detailed measurements of e.g. effects of the circadian clock on root growth in situ . To date, most circadian clock studies have been performed in the laboratory on young plants [38, 39]. Therefore, the necessity of developing a fully automated technique to measure root growth in the field is of major importance.
We compared the quality of images obtained, and the advantages or disadvantages when using several different types of scanner to measure root growth in the field. We focused on inexpensive technology that is currently available on the market and so is accessible to a wide range of potential users. We assessed these scanning techniques in conjunction with a fully manual method (tracing onto a plastic sheet) and a fully automated method (time-lapse camera). Measurements were performed in hybrid walnut (Juglans nigra × regia L.) agroforests in France. Results are discussed with regard to quality, time, and cost criteria.
Comparison of methods for acquiring images
We examined five methods for acquiring images of root systems growing in rhizotrons:
Scanners are lightweight (Fig. 1) and portable. We used a Vista Quest HS-500 (USA) to take images at 300 dpi. Scans were taken by moving the scanner manually downwards on the surface of the rhizotron window. Three A4 (21 cm wide and 45 cm long) images are needed for one 50 × 50 cm surface in order to include the borders of our rhizotrons (see section on “Rhizotron installation”). The images can be saved on a micro Secure Digital memory card and the scanner requires only two AA alkaline batteries to function.
If no electronic devices are available in the field, roots can be drawn manually with permanent color pens onto a transparent sheet placed over the rhizotron window. Colors indicate different observation times and the date of the observation is noted on the transparent sheet. Transparent sheets are then scanned in the laboratory using e.g. a scanner at a resolution of 300 dpi. (Sharp MX-3640 N PCL6, Canada). The manual tracing technique is not usually adequate for measuring root diameter precisely, because root diameter is not known, therefore it is not possible to select a pen with the appropriate point thickness. Nevertheless, manual tracing can be suitable for giving an estimate of root diameter class (e.g. Mao et al. ). In this study, we visually estimated root thickness and tried to use pens with the correct point thickness for tracing roots, so that we could compare results with those from the other methods.
Smartphone scanner application
Although often used to monitor the aerial phenology of vegetation, to our knowledge, time-lapse cameras have not yet been used for automated measurements of root growth and phenology in situ. Time-lapse cameras take photographs at regular intervals, determined by the user beforehand. We tested a Cuddeback Attack (U.S.A.) time-lapse camera with flash that takes photographs in color using LED bulbs (Fig. 1). Each camera was placed on a wooden cleat at a distance of 90 cm from the rhizotron. Photographs can be taken every 30 s (in our case, we took one photograph at 2 a.m. and at 12 h intervals thereafter). Time-lapse cameras can run for several months on an Alkaline battery (C (LR14) 1.5 V) without any human intervention.
Comparison of methods
Test 1: previously scanned and measured root systems
Test 2: Measurements in rhizotrons using scanners and manual tracing
We performed measurements at Le Beil, Madic, in the Cantal region, France (45°22′7.95″N, 2°28′1.46″E) (see section on study site for more details). We started the observations in October 2014 and fine root growth was measured every month from April to June 2015 using four methods: flatbed scanner, handheld scanner, smartphone scanner (iphone4) and the manual tracing (n = 25). Walnut fine roots are quite thick and so roots ≤ 4 mm in diameter were classed as fine roots.
Test 3: Measurements in rhizotrons using a time-lapse camera
A third set of measurements was performed at Cormont, in the Pas de Calais region, France (50°33′27.87″N, 1°44′3.08″E), (see section on study site for more details). Root growth was monitored in six rhizotrons twice a day from May to September. We focused our study on 21 roots growing over a period of 10 days for un easier understanding and comparison of results.
Once images of root growth had been acquired, we conducted analyses of images using the semi-automated SmartRoot software . SmartRoot is an operating system independent freeware based on ImageJ and uses cross-platform standards (RSML, SQL, and Java) for communication with data analysis softwares [36, 40]. Before analyzing roots with SmartRoot, when necessary, images need “stitching” together (e.g. with Adobe Photoshop CS3 software), if several have been taken (when the rhizotron surface area was greater than the field of the scanner). In our case, we transformed all images to 8 bit gray scale and then inverted them using ImageJ software so that roots were darker than the background of the image. The length and diameter of each root produced during one interval time (i.e. one month) were calculated for each rhizotron. Before analyzing a new sequence of images, SmartRoot provides the user with an icon to import the traces of the same roots from the previous image data file to superimpose them on this new image, which helps root elongation. This preceding image also helps determine whether the root is live (usually cream in color) or in a phase of senescence (shriveled, transparent or turning black) [3, 41–44]. We declared a root dead when it became completely black in color.
We measured fine root growth in situ in two agroforests. One was located at Le Beil, Madic, in the Cantal region, France (45°22′7.95″N, 2°28′1.46″E) at an elevation of 530 m, hereafter termed ‘continental’ climate. The agroforest comprised three transplanted tree species: hybrid walnut (Juglans major (MJ209) × Juglans regia L.), cherry (Prunus avium L.), sycamore maple (Acer pseudoplatanus L.) at 12 m × 8 m tree spacing and intercropped with permanent pasture (ovine or bovine pasture). All national guidelines and legislation were complied with when using these cultivars. Mean diameter at breast height (DBH) of all walnut trees at the site was 0.20 ± 0.02 m and mean height was 12.09 ± 1.30 m. Data are mean ± standard error. All trees were planted in 1994 at a density of 100 trees ha−1. Hybrid walnut at this study site starts leafing in early May and shedding in mid-November. The climate is continental with a mean annual temperature of 9.95 °C and a mean annual rainfall of 1174 mm (Météo France). The soil is silty and not deep, attaining an average maximum depth to bedrock of approximately 110 cm, on a 5°–10° slope.
The second agroforest was located at Cormont, in the Pas de Calais region, France (50°33′27.87″N, 1°44′3.08″E), hereafter termed ‘oceanic’ climate. The site is at an altitude of 40 m. The climate is oceanic, with a mean annual temperature of 11 °C and a mean annual rainfall of 777.9 mm (Météo France). Tree species comprised hybrid walnut (Juglans nigra × regia L.) and Maple (Acer laurinum L.) at 13 × 7.5 m tree spacing intercropped permanent pasture (ovine pasture). All trees were planted in 1999. The soil is silty clay and <2.5 m deep. The site is next to La Dordonne River. Mean DBH of walnut trees at the site was 0.30 ± 0.03 m and mean height was 14.75 ± 3.50 m. Hybrid walnut at this site starts leafing in early May and shedding in mid-November.
In the continental agroforest (Madic), we dug eight (1 m × 1 m × 1 m) trenches by hand in three rows of trees. Each trench was at a distance of 2 m from the nearest tree stem. Eight rhizotrons, or root windows (50 cm long × 50 cm wide × 0.5 cm thick), were installed. In the oceanic agroforest (Cormont), soil was deep (4 m) and comprised four (2 m long × 1 m wide × 2 m depth) trenches in one row of trees placed at 2 m from the nearest tree stem. One rhizotron was installed on two opposing faces of the trench (n = 8 rhizotrons in total). All rhizotrons were placed vertically at an angle of 15° from the face of the profile. This angle will permit the roots to grow downwards due to positive geotropism [21, 41]. Where the rhizotrons were to be placed on the trench, we gently removed the soil to make a flat surface and cut all roots on the profile with secateurs. The soil removed during the digging of the trenches was kept aside, and then sieved through a 5 mm size sieve and air-dried for several hours. The sieved and air-dried soil was then poured into the space between the window and the soil profile and slowly compacted using a wooden plank. Each rhizotron was covered with foil backed felt insulation and black plastic sheeting to protect roots from light and air temperature variations. All trenches were then covered with wooden boards and corrugated plastic to avoid damage from passing animals and to prevent direct rainfall and sunlight onto the rhizotrons. In the first three months after installation, no root growth was recorded to avoid over estimations of root growth .
Root indicator calculation
We used the following method to estimate root elongation rate: individual root growth was evaluated by calculating the difference between the root length at t−1 and at t. To determine the daily root elongation rate (RER), the mean of all individual root lengths produced between time t and t−1 was divided by the duration of the corresponding period . According to the literature, the characterization of dead roots is not obvious, particularly behind a transparent window . We considered a root as live when it had a cream color and dead when it had turned black with no growth between two or more successive sessions until the last observation date occurred .
Semi-quantitative scoring decision matrix
Three criteria were taken into account to evaluate the five methods: (i) accuracy (image quality and resolution, deformation and contrast), (ii) effectiveness (time, expenditure and labour) and (iii) adaptability (ease of use in field and necessity of accessories).
Root length and diameter obtained using each method were correlated with the previously scanned and measured root systems, to determine which method gave the best fit. Similarly, results from different generations of smartphones were compared. We then calculated relative values for cumulative length, mean diameter and RER, with regard to the flatbed scanner (reference value), which we assumed gave the most accurate dimensions [30, 32]. To calculate the relative value, we divided the value obtained for individual roots (using each method) by that obtained using the flatbed scanner.
A Shapiro–Wilk test was performed before each test to ensure if the investigated indicator followed a normal or non-normal distribution. Homogeneity of variances was checked. For data not normally distributed, analyses were followed by a Kruskal–Wallis Test for each factor. A post hoc analysis between root diameters was performed using the Nemenyi test of Kruskal–Wallis at p < 0.05 to determine which levels of the independent variable differ from every other level. All analyses were performed using R software, Version 2.15.3 (R Development Core Team 2013) at a significance level of <0.05.
Test 1: Previously scanned and measured root systems
Test 2: Measurements in rhizotrons using scanners and manual tracing
Test 3: Measurements in rhizotrons using a time-lapse camera
Studies on root growth have been numerous over the last decade and a significant progress in evaluating root morphology has been made. However, research remains challenging and costly especially in the natural environment. Many nondestructive methods, such as rhizotrons [18–21] and minirhizotrons [3, 22, 23] have been developed to overcome some of the limitations of observing root systems in the natural environment and to offer direct and repeated observations of root system morphology. Image quality obtained from rhizotrons and minirhizotrons is crucial for an accurate quantification of root growth through image analysis.
Multiple criteria evaluation/semi-quantitative scoring decision matrix
For 1 rhizotron
For 8 rhizotrons
Phone100–700€ Apps free
Usage in field
Yes/battery and PC
Time between visits
Time to treat images before analysis
To the best of our knowledge, the smartphone scanning application and an automated time-lapse camera have never been used to measure root growth in the field. Both methods are inexpensive and easy to use, especially compared to more sophisticated techniques such as minirhizotron scanners. The advantage of rhizotrons over minirhizotrons is that the above variety of inexpensive techniques exist worldwide for quantifying root growth in the field. The equipment needed to observe and record color video pictures of roots in minirhizotrons [9, 46] is commercially available at a cost of approximately 10,000 euros for one circular scanner  or one camera video. Additionally, the field of vision in a minirhizotron is small (20 × 20 cm) and is not suitable for heterogeneous forest stands, where the spatial position of roots of different diameter classes can lead to root-free patches in the soil. The 50 × 50 cm rhizotrons we used in our study enable more tree roots to be captured in one image, thus increasing statistical robustness.
We tested several methods for monitoring root growth and acquiring images in rhizotrons in the field. Our results show that scanners and time-lapse cameras provide correct measurements of root growth and length in the field but users should be aware of possible artifacts. Time-lapse cameras overestimate root diameter but are useful for taking frequent images of root elongation in the field over several months, without any manual intervention. Taking into account image accuracy, time spent and cost, we found the smartphone scanner to be the optimal method for monitoring root growth in the field. Future generations of smartphones could scan images and transfer data automatically, with a minimum of human intervention, thus improving the methodology. Likewise, the development of digital time-lapse cameras with a higher optical resolution, or similar to the optical resolution of smartphones, should also be undertaken.
root elongation rate
AM, YM and AS designed the study. AM, YM, AS, ZM and MR conducted fieldwork. AM, ZM and GL contributed to data analysis. AM and AS wrote the paper with contributions from YM, ZM, JLM and GL. All authors read and approved the final manuscript.
Thanks are due to François Pailler and Stéphane Fourtier (INRA) for technical assistance, to Camille Béral (Agroof) for help finding field sites, to Jérôme Nespoulous (INRA) for his help with field and laboratory work and to the farmers M. Queuille and M. Becue for letting us work in their agroforests.
Availability of data and materials
The datasets generated and/or analysed during the current study are available in the [Zenodo] repository, [https://doi.org/10.5281/zenodo.266959]
The authors declare that they have no competing interests.
Funding for a Ph.D. bursary was provided by Campus France and the Kurdish Institute, France (AM) and fieldwork was funded by the FOARDAPT project, INRA metaprogram AAFCC (Adaptation of Agriculture and Forests to Climate Change), France.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
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