Automated control offers the ability for continuous imaging and manipulation of media conditions with high temporal resolution. One notable example of a microfluidic device for rhizosphere studies is the RootChip, which uses the micro-valves in a PDMS device to control the fluidics . The first study using the RootChip grew 8 Arabidopsis plants on a single device with micro-valves but by the second iteration, the throughput has been doubled indicating rapid technological advances in the field. In addition, all these studies demonstrated spatiotemporal imaging at single-cell resolution and dynamic control of the abiotic environments in the rhizosphere. Another microfluidics-specific application to rhizosphere study is to use the laminar flow to generate the spatially precise and distinct microenvironment to a section of the root as demonstrated by Meier et al. . A young Arabidopsis’ seedling was sandwiched and clamped between two layers of PDMS slabs with microchannel features to tightly control synthetic plant hormone flow with 10 to 800 µm resolution to the root tip area, enabling observations of root tissues’ response to the hormones. As many root bacteria produce auxin to stimulate the interactions with the root, this study showed the possible mechanism of microbiome inducing the interaction by stimulating root hair growth. Another application of laminar flow utilized the RootChip architecture by adding the two flanking input channels to generate two co-laminar flows in the root chamber,hydroponic farming subjecting a root to two different environmental conditions along the axial direction to study root cells adaptation to the microenvironment at a local level . These studies revealed locally asymmetrical growth and gene pattern regulations in Arabidopsis root in response to different environmental stimuli.
Microfluidic platforms have also been successfully employed to study the interactions between the root, microbiome and nematodes in real time . In the systems, additional vertical side channels are connected perpendicularly to the main microchannel to enable introduction of microorganisms and solutes to the roots in a spatially and temporally defined manner . A recent microfluidic design incorporated a nano-porous interface which confines the root in place while enabling metabolite sampling from different parts of the root . These studies demonstrated the potential of microfluidics in achieving spatiotemporal insights into the complex interaction networks in the rhizosphere. Despite several advantages of microfluidics in rhizosphere research as described above, some challenges remain. All the microfluidic applications grow plants in hydroponic systems where clear media is necessary for the imaging applications and packing solid substrates in the micro-channels is not trivial. The microscale of the channels limits the applications of these devices to young seedlings. Thus, interrogating the microscale interactions in bigger, more developed plants is not possible with current microfluidic channel configurations. In addition, technical challenges such as operating the micro-valves and microfabrication present a barrier to device design and construction for non-specialists. Fabricated ecosystems aim to capture critical aspects of ecosystem dynamics within highly controlled laboratory environments . They hold promise in accelerating the translation of lab-based studies to field applications and advance science from correlative and observational insights to mechanistic understanding. Pilot scale enclosed ecosystem chambers such as EcoPODs, EcoTrons and EcoCELLs have been developed for such a purpose . These state-of-the-art systems offer the ability to manipulate many parameters such as temperature, humidity, gas composition, etc., to mimic field conditions and are equipped with multiple analytical instruments to link below ground rhizosphere processes to above ground observations and vice versa .
Currently, however, accessibility to such systems is low as there are only several places in the world which can host such multifaceted facilities due to the requirement of significant financial investments. Switching back to lab-scale systems, a recent perspective paper calls for the need to standardize devices, microbiomes and laboratory techniques to create model ecosystems to enable elucidation of molecular mechanisms mediating observed plant-microbe interactions e.g., exudate driven bacterial recruitment . Toward this goal, open source 3D printable chambers, termed Ecosystem Fabrication devices, have been released with detailed protocols to provide controlled laboratory habitats aimed at promoting mechanistic studies of plant-microbe interactions . Similar to a rhizotron setup, these flow through systems are designed to provide clear visual access to the rhizosphere with flexibility of use with either soil or liquid substrates . Certainly, there are many limitations to these devices in that they are limited to relatively small plants and limit the 3D architecture of the root system. Still, an advantage with the EcoFAB is that its 3D printable nature allows for adaptations and modifications to be made and shared on public data platforms such as Github for ease of standardization across different labs and experiments . In fact, a recent multilab effort showed high reproducibility of root physiological and morphological traits in EcoFAB-grown Brachypodium distachyon plants . The development of comparable datasets through the use of standardized systems is crucial to advancing our understanding of complex rhizosphere interactions. Open science programs such as the EcoFAB foster a transparent and collaborative network in an increasingly multidisciplinary scientific community. Specialized plant chamber systems are necessary for nondestructive visualization of rhizosphere processes and interactions as all destructive sampling approaches tend to overestimate the rhizosphere extent by 3–5 times compared to those based on visualization techniques . Nonetheless, plants in such chambers are still grown in defined boundaries and suffer from inherent container impacts.
For instance, studies have pointed out that container design significantly influences root growth during early developmental stages and leaves lasting impacts on plant health and phenotype . The majority of the lab-based chambers are also centimeter scale and are unlikely to replicate exact field conditions in terms of soil structure, water distribution, redox potential or root zone temperatures . While comparisons between chamber-grown and pot-grown plants show similar outputs , studies comparing plants grown in confined spaces to those directly grown in the field are missing. A recent review mapped the gradient boundaries for different rhizosphere aspects and found that despite the dynamic nature of each trait, the rhizosphere size and shape exist in a quasi-stationary state due to the opposing directions of their formation processes . The generalized rhizosphere boundaries were deducted to be within 0.5–4 mm for most rhizosphere processes except for gases which exceeds > 4 mm and interestingly, they are independent of plant type, root type, age or soil . Bearing this in mind, our assessment of the different growth chambers revealed possible overestimation of rhizosphere ranges in some chamber set ups. For instance, the use of root-free soil pouches representing rhizosphere soil despite being cm-distance away from the rhizoplane. This prompts the need for careful evaluation of new growth chamber designs to ensure accurate simulation of natural rhizosphere conditions. To date,ebb flow tray many rhizosphere microbiome studies and growth chambers systems focus on the impact of plant developmental stage, genotype and soil type on microbial composition and function . On the other hand, predation as a driver in the rhizosphere microbiome remains understudied. For instance, protists are abundant in the soil and are active consumers of bacteria and fungi and play a role in nutrient cycling yet remain an overlooked part of the rhizosphere . Viruses are also pivotal in modulating host communities thereby affecting biogeochemical cycles but their influence in the rhizosphere is poorly studied . These predatorprey interactions in the rhizosphere deserve in-depth studies which can be facilitated by these specialized growth chambers. Another area worth investigating in the rhizosphere is in anaerobic microbial ecology. At microbially relevant scales, soils primarily exist as aggregates . Aggregation creates conditions different from bulk soil, particularly in terms of oxygen diffusion and water flow resulting in anoxic spaces within aggregates and influences the microbial community . The rhizosphere is also rich in a wide range of compounds which can serve as alternative electron acceptors such as nitrate, iron, sulfate and humic substances in the absence of oxygen . However, most anaerobic studies in the rhizosphere focus only on aqueous environments such as water-logged paddy soils despite biochemical and metatranscriptomic evidence pointing to the possibility of anaerobic respiration in the rhizosphere . To fully understand biogeochemical cycles in the rhizosphere, it is imperative to investigate rhizosphere processes in the microscale and to include localized redox conditions as one of the influencing parameters. Microfluidic platforms with its fast prototyping capabilities can be helpful in creating growth chambers designed to stimulate these redox changes. In the study of the rhizosphere microbiome, genetic manipulation strategies are foundational in deep characterization of microbial mechanisms and current manipulation techniques require axenic isolates. However, the uncultivability of a significant portion of soil microorganisms continues to hamper efforts in gaining mechanistic knowledge. Even for culturable isolates, the process of isolation introduces selective pressure and disturbance to the community with inevitable loss of information on spatial interactions. A recent innovation in gene editing technologies using CRISPR-cas systems demonstrated in situ editing of genetically tractable bacteria within a complex community .
Coupled with the use of transparent soil-like substrates , the application of such a technique for the editing of in situ rhizosphere microbiome while preserving spatial and temporal associations would indeed bring invaluable insights. Specialized growth chambers using 3D fabrication and microfluidic technologies are primed to facilitate such innovations. Finally, this review revealed that while similarities exist among the different growth chamber systems, many of these systems are bespoke. This makes it difficult to replicate experiments and determine reproducibility which are important cornerstones of scientific advancement. The complexity of rhizosphere interactions also warrant that computational models are essential to gain a better understanding of system level processes . However, predictive modeling requires data from standardized approaches to be comparable between experiments. Thus, future growth chamber systems and designs are encouraged to follow the open science framework to enable standardization to an extent, such as in the case of EcoFABs . New root phenotyping technological developments are needed to overcome the limitations of traditional destructive root investigation methods, such as soil coring or “shovelomics” . Mancuso and Atkinson et al. provide extensive reviews on the methodological advances on non-destructive root phenotyping, including Bioelectrical Impedance Analysis , planar optodes, geophysical methods, and vibrating probe techniques. These techniques aim to mitigate key limitations of traditional root phenotyping, especially addressing the need for a better and more convenient characterization of the finer roots and root functioning. Advances in non-invasive and in-situ approaches for monitoring of root growth and function over time are needed to gain insight into the mechanisms underlying root development and response to environmental stressors. Geophysical methods have been tested to non-destructively image roots in the field. Ground Penetrating Radar approaches have been used to detect coarse roots . Electrical Resistivity Tomography and ElectroMagnetic Induction approaches have been used to image and monitor soil resistivity changes associated with the Root Water Uptake . Recent studies explored the use of multi-frequency Electrical Impedance Tomography to take advantage of the root polarizable nature . Despite these advantages, geophysical methods to date share common limitations regarding root characterization. Geophysical methods developed to investigate geological media: in the case of roots they measure the root response as part of the soil response, see Fig. 1a for the ERT acquisition. Because of the natural soil heterogeneity and variability the resolution and signal characteristics of geophysical methods strongly depend on soil type and conditions. As such, interpretation of the root soil system response is non-unique, hindering the differentiation between roots of close plants and the extraction of specific information about root physiology from the electrical signals. Unlike geophysical methods, the BIA for root investigation developed to specifically target the impedance of plant tissues, limiting the influence of the growing medium. A practical consequence is that BIA involves the application of sensors into the plant to enhance the method sensitivity. BIA measures the electrical impedance response of roots at a single frequency or over a range of frequencies . The measured BIA responses have been used to estimate root characteristics, such as root absorbing area and root mass . A key assumption is that current travels and distributes throughout the root system before exiting to the soil , with no leakage of current into the soil in the proximal root position . It is only in the former case, that the BIA signal would be sensitive to root physiology.