Application of Microfluidics in Plant Physiology and Development Studies

Abstract

Microfluidics has emerged as a powerful enabling technology in plant science, offering unprecedented control over microscale environments for the cultivation, manipulation, and analysis of plant cells, tissues, and organs. This review provides a comprehensive overview of the development and application of microfluidic systems in plant physiology and development studies. We categorize the platforms based on their structural designs and biological targets—from single-cell trapping devices and droplet-based screening systems to organ-on-a-chip and root–microbe interaction modules. Key applications include live-cell imaging, real-time monitoring of stress responses, microenvironment simulation, and high-throughput phenotyping. Particular attention is given to microfluidic investigations of plant mechanobiology, chemotropism, and cell-to-cell communication, as well as their integration with biosensors, electrophysiological tools, and environmental control systems. We also examine current limitations related to material compatibility, device scalability, and biological complexity, and highlight emerging solutions such as modular design, interdisciplinary integration, and soil-on-a-chip systems. By addressing both fundamental research needs and practical agricultural challenges, microfluidic technologies offer a transformative path toward precision plant science and sustainable crop innovation.

1. Introduction

Plant research encompasses multiple disciplines regarding usability for humans, such as ecology, biotechnology, agriculture, and medicine, each with its own goals and scales of study. While in situ field experiments capture natural interactions most authentically, they pose various methodological, environmental and logistical challenges while dissecting novel cellular pathways or traits: difficulty in continuous monitoring and sampling disturbances, restrictive permissions and regulations for transgenic organisms, uncontrollable biotic and abiotic factors, extensive land or greenhouse space and high resource demands [1,2,3]. To address some of these limitations, sensors and devices for live monitoring have been introduced in recent years [4]. However, standard approaches still primarily constitute of in vitro and in vivo cultivation of plant cells, tissues, organs or even whole plants grown in defined solid or liquid media (from <1 mL in multi-well plates to liters in bioreactors) [5,6,7,8] (Figure 1).

The three pillars of in vitro plant methods are: (i) callus and suspension culture on nutrient media, underpinning everything from secondary metabolite production to somatic embryogenesis [9,10]; (ii) micropropagation—rapid clonal multiplication via nodal segments, shoot tips or meristems [11,12]; (iii) organ culture, isolating shoots, roots or embryos to probe organ-specific physiology [13]. Media choice (solid vs. liquid) and vessel scale follow the biological question: small volumes for cell assays; larger jars or pots (often with soil) to mimic later developmental stages [14,15].

A critical control lever is the application of plant growth regulators (auxins, cytokinins, gibberellins) to steer morphogenesis—callus induction, shoot vs. root formation, etc. [9,10,16]. Environmental synchronization (light cycles, temperature shifts, osmotic stress) further allows cell-cycle studies or developmental gating [17,18].

Beyond basic culture, specialized techniques include genetic transformation (Agrobacterium-mediated, biolistics) for functional genomics [19,20] and cryopreservation of germplasm for long-term storage and biodiversity conservation [21,22].

These established methods power everything from crop improvement to metabolic engineering. But to tackle future challenges, such as screening thousands of genotypes and dynamically probing single-cell responses, miniaturization and parallelization are key, which can be strongly promoted by the application of microfluidics.

Microfluidic technology has transformed experimental biology by enabling precise manipulation of fluids at the nanoliter to picoliter scale. By miniaturizing assay environments, microfluidics drastically reduces the consumption of reagents and biological samples, making experiments more cost-effective and resource-efficient [23,24]. The small scale of these systems accelerates processes through rapid diffusion and enables automated control over fluid flow [24]. One of the most impactful features of microfluidics is its capacity for high-throughput analysis: multiple assays or reactions can be performed simultaneously on a single chip, significantly increasing experimental throughput. Moreover, microfluidic platforms can integrate various laboratory steps, such as sample preparation, reaction execution, and detection, into one compact system, which not only streamlines workflows but also minimizes contamination and manual intervention [25]. In parallel, simplified microfluidic strategies have been developed to reduce operational complexity, including passive and pumpless approaches that rely on capillary forces, surface wetting, or evaporation-driven flow. By minimizing the need for external pumps and tubing, such systems lower technical barriers and may facilitate broader adoption of microfluidics in long-term and parallelized biological studies [26].

While most advances have targeted microbial or mammalian cells, plant biology presents unique demands (cell wall, larger cell size, photosynthetic requirements)—each calling for tailored channel geometries, material choices and light delivery strategies [27].

In this review, we will survey plant-focused microfluidic platforms used for cultivation and analysis in plant research, along with current trends and directions in their development.

2. Plant Cells, Tissues, and Organs in Microenvironments

2.1. Plant Research Fields with Advantageous Application of Microfluidic Devices

Microfluidic systems designed for the cultivation and analysis of plant tissues and organs typically consist of microchannels, chambers, and integrated structures that permit continuous microscopic observation during growth, using light or fluorescence microscopy. These devices are most commonly fabricated from polydimethylsiloxane (PDMS), often bonded to glass substrates to enhance optical clarity. PDMS is widely used due to its biocompatibility, gas permeability, and ease of prototyping, although it may present limitations in certain imaging modalities or chemical interactions. The complexity of plant-oriented microfluidic platforms varies, ranging from simple channel networks that guide tissue growth to advanced architectures tailored for specific experimental goals, such as chemical gradient generation, root–microbe interaction studies, or hormone transport analysis. A broad and growing spectrum of applications has been explored using such systems.

Table 1. Microfluidic devices for plant research applications.

Plant Research Application Field	Microfluidic Device/Platform	Cell Type/Tissue/Organ of Interest	Refs.
Cell-based approaches
Single cell and protoplast analysis	Chamber-based microfluidic device	Protoplasts, protonema cells, somatic cells	[28,29,30,31,32]
	Single cell trapping devices	Protoplasts, pollen tubes, ovules	[33,34,35,36,37,38]
	Microwells array	Protoplasts	[39,40]
	Size-based microarrays	Protoplasts, chloroplasts	[41,42]
	Channel-based microfluidic devices	Protoplasts, pollen tubes, somatic cells, root hairs	[43,44,45,46,47,48,49,50,51,52]
	Droplet-based microfluidics	Protoplasts, microspores	[53,54,55,56,57,58]
	Impedance-based microfluidic devices	Protoplasts, mesophyll cells, microspores, pollens	[59,60,61,62,63,64,65]
	Microfluidic devices for cell stretching and compression	Pollen tubes	[66,67,68]
Development and structural characterization
Morphological/functional phenotyping	Chamber-based microfluidic device	Protonema tissue	[69]
	Microfluidic chip arrays	Germinating seeds	[70,71,72,73]
	Channel-based microfluidic devices	Root hairs, roots	[74,75]
	Microfluidic devices coupled with sampling fluidics	Roots	[76]
	Root array	Roots	[77]
	Root-on-a-chip device	Roots	[78,79,80,81,82,83]
	Rhizosphere-on-a-chip device	Roots	[84]
	Microfluidic capillary zone electrophoresis	Root extracts	[85]
Environmental interactions and responses
Abiotic stress response	Root-on-a-chip devices	Roots	[86,87]
	Microfluidic devices for precise control of environmental conditions	Roots	[88]
	Microchannel-based microfluidic devices	Roots, plant extracts	[89,90,91,92,93,94]
	Chamber-based microfluidic devices	Roots, leaves	[95,96]
	Microfluidic chip arrays	Roots	[97]
	Microgravity generating devices	Seeds	[98]
	Rhizosphere-on-a-chip devices	Roots	[99]
	Paper-based sensors	Plant extracts	[100,101]
	Microfluidic-printed electro-mechanical sensors	Leaves	[102]
Cell communication and plant-microbe interactions	Modular microfluidic bioreactors	Somatic cells	[103]
	Trapping devices	Somatic cells, roots	[104]
	Microfluidic chambers for co-cultivation	Roots	[105,106,107,108]
	Soil-analog microfluidic devices	Roots	[84,109]
	Microchannel-based devices	Pollen tubes, roots	[110,111,112,113,114,115]
	Bioelectricity generating devices	Roots exudates	[116]
	Microfluidic-based single-nucleus RNA-seq	Root extracts	[117]

provides an overview of representative microfluidic platforms, 2m2m2,1,111categorized by their application area in plant research and the associated plant tissues or organs involved.

Some fields of applications particularly benefit from utilization of microfluidics throughout the process of investigation when compared to the standard methods of plant cultivation and analysis. Microdevices can be manufactured to either support or constrain the growth of specific plant organs, depending on the experimental aim. Additional microstructures assist in the immobilization, shaping, and positioning of samples, thereby enabling visualization and tracking of the target tissues or cells. Control of the media composition and creation of constant fluid flow allows for real-time observation of the effects and dynamics of environmental conditions on the objects of interest. The inclusion of sensors into the system design gives valuable information about the physical or chemical state of the cells and tissues.

In the next chapters of this review, we present a detailed overview of the main groups of aforementioned microfluidic devices, focusing on their current applications, limitations, and potential directions for further development.

2.2. Microchambers and Microchannels in Plant Cultivation

Although plant systems introduce unique biological requirements, such as rigid cell walls, larger cell sizes, and complex tissue organization, many core microfluidic strategies, originally developed for animal cells, microbes and algae, are transferable, provided appropriate design modifications are made [118,119].

When engineering microfluidic platforms for plant cultivation, several critical factors must be addressed:

• Fluid flow and shear stress: Plant cells vary significantly in their sensitivity to mechanical forces; for example, fragile root hairs and protoplasts require gentle flow regimes to prevent structural damage [120,121].
• Gas exchange: Unlike some animal cell cultures, plant systems depend on efficient CO₂ and O₂ diffusion, particularly during photosynthesis and root respiration [122,123].
• Cultivation volume: Optimal chamber or channel volume must balance sufficient nutrient delivery with the need to maintain biologically relevant interactions such as quorum sensing or paracrine signaling [124].

Microchambers represent a foundational component of plant-on-chip technologies, offering precise spatial and temporal regulation of environmental conditions. Their confined volumes—typically ranging from nanoliters to microliters—are particularly well-suited for high-throughput cultivation of single cells or organoids under controlled conditions. In contrast, larger cultivation chambers are the default choice for studies involving seeds, roots, and shoots. Depending on the plant species and developmental stage, these chambers often extend into the millimeter or even centimeter scale.

As illustrated in Figure 2, microchambers exhibit diverse architectural configurations tailored to specific experimental needs. For example:

(a)

A dual-layer design separates a thin cell-cultivation chamber from an underlying transfer channel via a porous membrane, enabling continuous metabolite exchange [31].

(b)

A pentagonal array of interconnected chambers (~160 nL each) permits parallel cultivation of plant protoplast populations, with integrated microcolumn structures (20 μm gaps) that prevent cellular escape [29].

(c)

Large-scale arrays composed of square wells facilitate seedling cultivation and phenotyping; these systems support simultaneous exposure to multiple media compositions and allow for high-throughput screening [72].

(d)

Whole-plant or organ-level cultivation is enabled by macro-scale chambers (cultivation areas > 85 cm²), such as the Root-TRAPR system, which accommodates expansive root architectures while maintaining optical access for imaging and analysis [78].

These platforms ensure high control demonstrated on biologically relevant examples, and they are instrumental in plant tissue culture and morphogenesis studies.

Microchannels serve a complementary function by enabling guided growth and manipulation of single cells, as well as multicellular structures such as roots, pollen tubes, and vascular bundles. Their linear or branched geometries provide laminar flow for nutrient delivery and chemical gradient formation, directional guidance for elongating structures (e.g., root hairs, pollen tubes), as well as access for real-time, non-invasive imaging and spectroscopic analysis. Depending on the scientific aim and the size of investigated structure, channels vary in their diameter from just 1 μm (as in the investigation of cell abilities to pass through extremely narrow spaces [47]) up to 800 μm for accommodating tree roots early growth [111].

Figure 3 showcases a variety of channel-based applications: directional pollen tube growth (Figure 3a) [45], single-cell analysis via flow cytometry (Figure 3b) [59], cell nuclei in root hairs localization (Figure 3c) [75], and dual-flow configurations for asymmetric stimulus exposure (Figure 3d) [87]. These devices provide valuable insight into developmental and physiological processes with fine spatial resolution.

In parallel with these architectural developments, alternative fabrication strategies are increasingly being explored to improve accessibility, scalability, and customization of plant microfluidic platforms. In particular, three-dimensional (3D) printing has emerged as a complementary approach to conventional soft lithography for the fabrication of microchamber- and channel-based systems. Several of the platforms discussed above demonstrate the feasibility of additively manufactured components across different biological scales. Ecosystem fabrication (EcoFAB) devices employ widely available 3D printing technologies to construct modular growth environments for studying plant–microbe interactions under controlled conditions [107]. Similarly, the Root-TRAPR system integrates a 3D-printed structural frame with optically transparent components to enable reusable, low-cost cultivation of larger plant root systems [78]. At the microscale, stereolithography-based 3D printing has been successfully applied to fabricate open microfluidic root imaging platforms, while careful material selection remains essential to ensure plant biocompatibility [86]. Beyond cultivation systems, recent work combining 3D-printed microfluidic transfer tools with droplet-based workflows has enabled image-guided handling and sorting of plant-derived cells such as pollen, highlighting the compatibility of additive manufacturing with advanced single-cell microfluidic operations [125]. Together, these examples illustrate how 3D printing can support rapid prototyping, modular design, and adaptation of microfluidic devices to diverse plant research applications.

Although many microfluidic platforms are optimized for cellular- or tissue-level investigations, an important consideration is the extent to which insights obtained under microscale confinement can be extrapolated to whole-plant physiology. The highly controlled environments characteristic of microfluidic systems enable precise dissection of localized responses, such as mechanosensing, chemotropism, or cell-to-cell signaling; however, whole plants integrate these responses across multiple organs and spatial scales. Consequently, direct quantitative translation from microfluidic conditions to macroscopic growth behavior remains challenging. Nevertheless, microfluidic studies provide valuable mechanistic understanding that can inform plant-level hypotheses when interpreted in the context of system-level regulation. Emerging strategies to bridge this scale gap include modular device architectures, root-focused platforms that preserve shoot development ex situ, and mesofluidic or ecosystem-inspired systems that accommodate larger cultivation volumes while retaining environmental control. Together, these approaches position microfluidics not as a replacement for whole-plant experiments, but as a complementary tool that links cellular mechanisms to organism-level function.

In the following chapters, we will explore specific microfluidic devices based on microchambers and microchannels, highlighting their applications in plant cell manipulation, tissue culture, and organ-level investigations.

3. Microfluidically Assisted Plant Technology

The potential applications of microfluidics in plant research are diverse. In precision agriculture, microfluidic systems allow for localized delivery of nutrients or pH adjustment directly at the root zone, optimizing growth conditions and reducing resource waste. In hydroponic and soilless cultivation, microfluidic technologies provide precise control over nutrient supply and root-zone environments, offering significant advantages in regions with limited arable land. Furthermore, integration of microfluidic-based biosensors enables real-time monitoring of plant exudates and biochemical markers, facilitating the early detection of plant diseases and supporting timely intervention strategies. Microfluidic systems can also be incorporated into controlled environment agriculture, such as vertical farming and greenhouse cultivation, where dynamic adjustment of environmental parameters can support the development of more resilient crop varieties.

In the following subchapters, we review recent advances in microfluidic applications for plant research, structured into three major areas: (1) cell-based investigations, (2) studies of plant development and structural characterization, and (3) investigations into plant cell stress responses and intercellular communication.

3.1. Microfluidic Devices for Cell-Based Investigation

Microfluidics technology, when operating in micro and nano dimensions, offers unique opportunity for the investigation of single cells, which is difficult to obtain using conventional research methods. Traditional approaches typically yield data averaged across entire cell populations, masking the variability and individual behaviors of cells. In contrast, single-cell or near-single-cell studies provide detailed insights into the specific physiological and molecular states of individual cells. To enable such investigations, microfluidic devices must meet specific criteria in terms of scale, fluid control precision and shear stress minimization. Given the broad range of studies in this area, we categorize microfluidic platforms into four main types, based on their structural and functional features: (1) chamber- and microstructure-based systems, (2) microchannel-based systems, (3) microsystems for cell sorting, and (4) systems designed to investigate plant cell biomechanics.

We begin by presenting key examples of chamber-based and structured microenvironments developed for plant cell studies. These devices typically employ confined wells, traps, or microfabricated cages to immobilize single cells or tissues for extended monitoring, cell fusion studies, or developmental analyses. The following studies illustrate the diversity of such platforms in terms of their technical features and biological applications (

Table 2. Chamber- and microstructures-based systems for plant cell investigation.

Device Type	Purpose	Characteristics	Cell Type, Organism	Ref.
Culturing chambers array	Protoplasts long-term cultivation and imaging	Array of 36 PDMS culturing chambers (diameter of 4 mm, depth of 500 µm) covered with glass or PDMS covers, placed in a square dish	Protoplasts, Nicotiana tabacum	[28]
Microchamber-based pentagonal array	Protoplasts fusion, cultivation and imaging	Five culture chambers (900 µm in width, 3200 µm in length and 55 µm) arranged in a pentagonal array, each containing Double micro-column (30 µm in length, 20 µm in width, and 55 µm in height) line for protoplasts trapping	Protoplasts, Nicotiana tabacum	[29]
U-shaped trapping microfluidic platform	Cultivation and microscopical imaging	Array of 112 flow-through U-shaped traps (inner length 60 µm, inner width 45 µm), arranged in 14 lines	Protoplasts and spores, Physcomitrella patens	[33]
Shallow microfluidic chamber	Protonema culture and real-time cytoskeleton imaging under highly inclined and laminated optical sheet microscopy (HILO)	Cultivation chamber (2 mm wide, 12 mm long, with height varying between 4.5 and 15 µm), containing supporting pillars	Protonema cells, Physcomitrella patens	[30]
Microfluidic antibody capture (MAC) chip	Single cell protein expression quantification	50 analysis chambers (volume of 0.75 or 4.5 nL) with a micro-printed antibody spot individually connected to one reservoir channel used for cell and solutions delivery	Protoplasts, Arabidopsis thaliana	[32]
Microfluidic chip for single and double cell trapping	Controlled induction and study of protoplast fusion dynamics	Channel height of 56 µm, trapping features of minimum width 90 µm, consisting of 20 or 40 µm diameter posts arranged in a double U-shape.	Protoplasts, Arabidopsis thaliana	[34]
Microfluidic chip with an orifice array	Electrofusion of protoplasts	Two protoplast chambers between glass electrodes (425 µm spacing), separated by a Kapton sheet with a 5 µm-diameter, 25 µm-thick orifice array	Protoplasts, Phalaenopsis, Raphanus raphanistrum subsp. sativus	[36]
Microwells array	Shaping the cells in controlled geometries, microscopical imaging	Microchambers of circular, triangular, square and rectangular shapes with diameter of 15–40 µm and height 20 µm	Protoplasts, Arabidopsis thaliana	[39]
Microwells array	Cortical tension generation via cells confinement and cortical microtubules (CMTs) imaging	Microwells of various dimensions: 15 × 20 µm, 14 × 14 µm and 12 × 40 µm used for protoplasts confinement in various osmotic pressure conditions (280, 600 or 800 mOsmol·L−1)	Protoplasts, Arabidopsis thaliana	[40]
Microvessels array	Trapping and microscopical imaging	Chamber consisting of microvessels 72.8 µm × 52 µm	Protoplasts, Nicotiana tabacum	[35]
Microcage array	Immobilization and cultivation of ovules	Array of microcages of 650 µm length and varying width: 150, 200, 250 and 300 µm with PDMS pillars surrounding trapped ovule, for long-term ovule culture 200 and 250 µm wide microcages were used	Ovules, Arabidopsis thaliana	[37]

).

A wide range of chamber designs has been developed for plant cell cultivation and observation, reflecting the diverse functional demands of microfluidic assays—from maintaining cell viability to enabling controlled interactions or fusion events. One example emphasizing physiological support rather than mechanical manipulation is the PDMS-based microculture system introduced by Ju et al., who demonstrated that using an air-permeable PDMS cover sheet of optimized thickness (400 µm) significantly enhances oxygen delivery, enabling long-term protoplast culture and microcolony formation over 10 days [28]. Beyond cultivation, chambers can also function as biochemical assay units. Mickleburgh et al. developed a miniaturized microfluidic antibody-capture chip featuring dozens of nanoliter-scale chambers, each containing a printed antibody spot, enabling single-protoplast isolation, optical–chemical lysis, and quantitative single-molecule protein detection [32].

Depending on the experimental objective, chambers may also incorporate microstructures for cell immobilization or controlled pairing. For instance, protoplast fusion can be studied in relatively large chambers where cells interact stochastically and a simple microcolumn line provides minimal trapping [29], or alternatively in microdevices featuring arrays of U-shaped microstructures designed to hold a maximum of two protoplasts at a time [33]. Similarly to the latter approach, Seidel et al. presented a microfluidic device in which the observation chamber is covered with H-shaped trapping features formed by arranged posts. In their system, fluids can travel through the array in two directions, allowing for better placement and removal of double-trapped protoplasts [34]. In addition to these mechanically based trapping strategies, Hung et al. developed an electrofusion platform that uses an insulating orifice sheet between two Indium Tin Oxide (ITO) electrodes to align protoplasts by positive dielectrophoresis and achieve one-to-one fusion through pulsed electric fields [36].

For certain cells, such as the protonema cells of the moss Physcomitrella patens, trapping is not required. Instead, supporting pillars can serve as anchoring points, allowing the investigation of individual cells within a shared, large (2 × 12 mm), but thin (4.5–15 µm height) chamber filled with small clusters of cell via tubing [30]. The same organism was also studied using the aforementioned U-shaped trapping array (Figure 4a) [33]. Enclosing cells in confined spaces can serve not only as a trapping method but also as a means to investigate changes in cell geometry and cytoskeletal organization. Microdevices designed for such purposes often take the form of arrays composed of microvessels (Figure 4c) [35] or microwells (Figure 4d) [39]. Colin et al. further used microwell confinement to show that cortical tension, rather than geometry alone, governs microtubule orientation in protoplasts, with increased pressurization inducing a switch from longitudinal to transverse alignment [40]. This trapping approach can also be applied to more complex plant structures, such as ovules. A key design consideration in this case is the significantly larger dimensions required for the trapping microstructures—250 µm wide and 650 µm long cages allowed for successful cultivation of single ovules for long-term imaging.

The variety of trapping devices alongside their dimensions are presented in Figure 4.

While chamber- and structure-based systems excel in immobilizing plant cells for prolonged observation or precise manipulation, other microfluidic approaches aim to recreate flow dynamics, gradient conditions or accommodate cell growth. Microchannel-based platforms are briefly described and summarized in

Table 3. Microchannel-based devices for plant cell investigation.

Device Type	Purpose	Characteristics	Cell Type, Organism	Ref.
Microfluidic chip with integrated concave and convex–concave microsieve arrays	Protoplasts collection and lysis	Main flow channel (600 µm width) and a protoplast sieving array (with a square length of 25 or 50 µm and distance between microsieves of 10 µm) with collecting channels (300 µm width)	Protoplasts, Phalaenopsis Chiada Pioneer	[41]
Deterministic lateral displacement (DLD) arrays integrated into one microfluidic chip	Size-based Chloroplast separation	Four parallel DLD arrays with 10-µm pillars and gap spacings from 5 to 11 µm, each providing a distinct critical diameter and enabling simultaneous size-based separation of chloroplasts in the 2–5 µm range	Chloroplasts, Spinacia oleracea L.	[42]
Microchannel with hydrodynamic trap	Analysis—electrical permeabilization for extracting cytosolic molecules	Microchannel with hydrodynamic trap for clusters of intact plant cells analysis	Protoplasts, Arabidopsis thaliana	[43]
Microchannel based device	Pollen tube guidance and cultivation	Multiple microchannels 5–20 µm wide	Pollen tubes, Torenia fournieri	[44]
TipChip— microfluidic network	Pollen tube guidance and cultivation	Network of microchannels 30 µm wide and 80 µm deep for pollen tube growth with additional structure elements: kink for locking the pollen grain, sections of air-media interface and additional inlet for creating chemical gradient	Pollen tubes, Camelia japonica	[45]
Microfluidic network based on the TipChip	Real-time Manipulation and analysis of pollen tube growth direction	Planar fluidic network with a depth of 80 µm, consisting of a main chamber into which the pollen grains are injected, two traps with adjacent microchannels into which the pollen tubes elongate, and two side inlets for the injection of different media into the main chamber	Pollen tubes, Camelia japonica	[46]
Microchannel-based device	Cell cultivation through extremely narrowed spaces	Growth channels for tip growing cells with microgaps of 1 or 4 µm width	Pollen tubes, T. fournieri Root hairs, A. thaliana Somatic cells, P. patens	[47]
Microchannel-based device	Cell guidance and testing of cell wall rupture	500 µm long channels with narrowed gaps of various wide-length parameters: 4–20, 7–20 and 4–400	Pollen tubes, Arabidopsis thaliana	[48]
Microchannel-based device	Protoplasts cultivation and imaging	Main microfluidic channel (13.8 mm long, 0.1 mm high and 1 mm wide) with oval region for current reduction, posts for decreasing of the shear stress and columns line (12 µm gaps between each column) for trapping protoplasts	Protoplasts, Nicotiana tabacum	[49]
Microchannel-based sorting chip	Cell sorting	100-µm-wide, 80-µm-deep channels guide cell flow; sorting is triggered by bright-field and/or fluorescence-based classification, directing cells into either the sorted or unsorted outlet	Protoplasts, Arabidopsis thaliana	[50]
Crossed microfluidic chip	Cell wall regeneration, protoplasts cultivation and imaging	Three crossed microfluidic channels with a width of 250 µm, depth of 60 µm, and length of 30 mm (5–6 mm crossed zone) per device, 4 devices on a chip	Protoplasts, Nicotiana tabacum	[51]
Lab-on-a-chip device	Pollen grain trapping, pollen tube cultivation	Microfluidic network with hydrodynamic trapping of pollen grains with growth microchannels of various shapes	Pollen tubes, Camilla japonica	[38]
Droplet-based microfluidic device	Protoplasts cultivation	Top plate of the channel equipped with magnet, allowing for lining of magnetic microparticles labeled protoplasts for visualization	Protoplasts, Arabidopsis thaliana	[53]
Droplet-based microfluidic device	Protoplasts handling and fluorescence measurement analysis + sorting	Flow-focusing generation of droplets with protoplasts encapsulation, followed by on-chip fluorescence measurements	Protoplasts, Marchantia polymorpha	[54]
Microdroplet generation system	Protoplasts handling and cultivation	Agarose droplets generated via droplet chip junction	Protoplasts, Nicotiana tabacum	[55]
Microdroplet generation system	Identification and development of embryogenic microspores	Microspores individually introduced to the required volume transmitted via tubing to the tip of the capillary (final droplet volume 35–100 nL, coated by 800–1000 nL of mineral oil	Microspores, Hordeum vulgare	[56]
Droplet-based microfluidics	Induction and optimization of embryogenic development in microspores	Droplets of volume around 120 nL generated using 6-channel droplet generator and cultivated in polytetrafluoroethylene (PTFE) tubing of 0.5 mm inner diameter, separated from each other by perfluoromethyldecalin (PP9)	Microspores, Brassica napus	[57]
Droplet-based microfluidics	Protoplasts cultivation	Droplets of volume around 300 nL generated using 6-channel droplet generator and cultivated in PTFE tubing of 0.5 mm inner diameter, separated from each other by PP9	Protoplasts, N. tabacum, Brassica juncea, Kalanchoe daigremontiana	[58]

.

Depending on the target application, microfluidic channels for plant cell studies vary in size. They can be relatively large, serving as transport pathways or chamber analogs, or just slightly wider than the cell diameter to direct growth. Protoplasts, for example, fall into the former category. Microfluidic devices for protoplast research typically focus on isolation, lysis, cultivation, or analysis. A distinctive example is a device featuring multiple U-shaped microchannels with concave and convex-concave sieving arrays, enabling rapid protoplast extraction, collection, and lysis [41]. A related organelle-scale approach employs parallel deterministic lateral displacement (DLD) arrays for size-based chloroplast isolation, demonstrating that similar sieving principles can be adapted to subcellular targets with appropriate geometric scaling [42]. Cultivating protoplasts in large channels while maintaining fluid flow exposes them to mechanical stress, which is detrimental to these wall-less, fragile cells. To address this, Ko et al. designed a device incorporating an oval region to reduce flow velocity, posts of varying sizes to lower shear stress, and two column lines for protoplast trapping [49]. Notably, simpler crossed-channel systems have also been shown effective for processes such as cell wall regeneration, without the need for additional structures [51].

Droplet-based microfluidics offers another route for single protoplast observation. Large droplets (~4.5 µL) containing multiple protoplasts labeled with magnetic microparticles can be aligned using a magnet-equipped top plate for controlled evaluation [53]. To encapsulate individual cells, smaller channels and precise techniques such as cross-focusing droplet formation are required, enabling high-throughput generation and sorting of single-cell droplets [54,55].

Beyond hydrodynamic approaches, Landenberger et al. demonstrated an active optical-sorting strategy in which dynamically steerable optical tweezers displace and sort individual biological particles, including 30-µm plant protoplasts, based on morphology or fluorescence signals within a microfluidic chamber. This platform highlights the compatibility of optical gradient forces with plant cell handling and offers a flexible, label-adaptive alternative to geometry- or flow-based sorting methods [50].

Pollen tubes, among the fastest-growing plant cells, are ideal targets for microchannel-based investigations. Horade et al. developed a microchannel array for pollen tube growth, concluding that channels measuring 5–12 µm in both height and width support the insertion of single, intact tubes [44]. Around the same time, Agudelo et al. introduced TipChip, a modular microelectromechanical systems (MEMS)-based platform with a growth chamber, hydrodynamic traps for pollen grains, and microchannels of varying geometry to study tube growth, reorientation, and responses to air or chemical gradients (Figure 4a) [38,45,46]. Yanagisawa et al. further demonstrated that pollen tubes, root hairs, and moss protonemata can elongate through gaps as narrow as 1 µm [47], while other studies have used channel geometry modifications, in particular narrowing of channels (4–7 µm in diameter) for various length (20 or 400 µm), to investigate protein contributions to cell integrity under mechanical constraints [48].

Beyond protoplasts, other plant cell objects such as microspores are promising candidates for application of droplet-based microfluidics. In androgenesis research—where efficiency is typically low—creating numerous, individually monitored microenvironments is advantageous. Early demonstrations of droplet generation via microinjection date back to the 1990s [56], and more recent work shows embryogenic development of microspores in droplet- and tube-based system with volumes around 130 nL, allowing systematic screening of stimuli and effectors [57]. This platform has since been further developed to accommodate additional plant cell types, and Marczakiewicz-Perera et al. have demonstrated its applicability for prolonged protoplast cultivation and nearly single-cell–level observation [58].

Microchannels are also widely used for analytical applications. Cells can be transported to defined locations for targeted analysis, as in the case of RNA extraction using focused electric fields. Here, for example, intact cell clusters travel through a 100 µm deep and wide channel to a 3 µm hydrodynamic trap, facilitating molecular extraction for downstream analysis [43]. Additional microfluidic platforms for cell characterization and analysis are summarized in

Table 4. Analytical microsystems for plant cell characterization.

Device Type	Purpose	Characteristics	Cell Type, Organism	Ref.
Microfluidic chip-electrochemical detection (ED) system	Detection of ascorbic acid in single cells	Device consists of sample, separation and waste channels, each with a depth of 30 µm and a width of 80 µm, arranged in the double T-injector. Protoplasts are separated, lysed and analyzed using changing voltage (100–50 V for injection, 1000 V for lysis and 0.90 V for AA detection)	Protoplasts, Triticum aestivum	[52]
Microfluidic flow cytometer	Analysis—fluorescence detection Auxins level during cell wall regeneration Cytosolic redox status	Flow-through channel 60 µm high and 40 µm wide, accommodating single cells	Protoplasts, Arabidopsis thaliana, Petunia	[59]
Microfluidic impedance spectroscopy platform	Analysis—impedance measurement characterization of single cell at different plant cell wall regeneration	Platform contains a sensing trap and reference trap (which remains empty for the time of measurement)	Mesophyll cells, Arabidopsis thaliana	[60]
Microfluidic flow cytometry platform with fluorescent, mechanical and electrical modules	Analysis—fluorescence microscopy and impedance measurement	Platform contains flow-through channel accommodating single cells, equipped in electrodes for electrical-mechanical detection	Protoplasts, Arabidopsis Columbia	[61]
Microfluidic-based impedance flow cytometry platform	Analysis—impedance measurement at the frequency 0.5 to 12 MHz		Immature and mature pollen grains, Nicotiana tabacum (microscpores), Cucumis sativus, Capsicum anuum, Solanum lycopersicum	[62]
	Simultaneous mechanical and electrical characterization, At the frequency 0.5 and 5 MHz		Protoplasts, Arabidopsis thaliana, Populus trichocarpa	[63]
Microfluidic-based impedance flow cytometry platform	Cell viability measurements, detection of different microspore developmental stages during pollen formation and androgenesis	The Coulter system with coupled microfluidic chips containing electrodes and a microchannel of various sizes	Microspores, Triticum aestivum	[64]
Microfluidic flow cytometry platform	Characterization of cells subpopulations based on the biophysical properties using direct-current insulator-based dielectrophoresis (DC-iDEP)	Platform contains flow-through channel accommodating single cells, equipped in electrodes for electrical-mechanical detection	Protoplasts, Lophatherum gracile Brongn, Phyllostachys heterocycle ‘Pubescens’	[65]

.

Microfluidic-based analytical tools for single-cell investigations often adapt standard bulk-culture techniques to smaller sample volumes, offering enhanced sensitivity and dynamic monitoring capabilities. These systems, including microfluidic flow cytometers and impedance spectroscopy platforms, are crucial for analyzing plant cell metabolic activity, viability, and developmental fate at the individual cell level. Among the early microfluidic analytical approaches, Xia et al. introduced a single-cell electrochemical detection platform integrating cell injection, electrokinetic trapping, electrical lysis, and end-channel amperometric readout within a double-T microfluidic architecture. Their system enabled quantification of intracellular ascorbic acid in individual Triticum aestivum protoplasts, demonstrating how microfluidics can couple precise cell handling with sensitive electrochemical detection to analyze electroactive metabolites at the single-cell level [52].

Microfluidic flow cytometry enables high-throughput single-cell analysis by transporting cells through narrow microchannels that fit individual cells, thus facilitating the assessment of population heterogeneity. Dai et al. developed a microfluidic flow cytometry system based on fluorescence detection, enabling single-cell measurements of intracellular reactive oxygen species (ROS) levels under various environmental stress conditions. Their study also demonstrated the photoprotective role of anthocyanins and the promotion of primary cell wall (PCW) formation by auxin at the single-cell level, using protoplasts as the model system [59].

Label-free impedance flow cytometry (IFC) platforms offer an alternative approach for characterizing plant cells by measuring their mechanical and electrical properties. Han et al. incorporated a constriction channel into a microfluidic IFC platform, allowing differentiation of protoplasts derived from different species and transgenic mutants overexpressing specific plasma membrane proteins. The two key parameters measured, passage time and impedance opacity, provided information on cell deformability and membrane electrical properties, respectively [63]. IFC was also successfully employed to assess pollen viability and germination potential. Heidmann et al. demonstrated that IFC could distinguish between dead and viable pollen based on cell size (with dead cells exhibiting reduced size), membrane integrity, and cytoplasmic conductivity across several species [62]. Similarly, Canonge et al. applied IFC to wheat microspores to monitor gametophytic and sporophytic developmental pathways, enabling early prediction of embryo yields and providing a robust marker for androgenesis efficiency [64]. Liu et al. further showed that dielectrophoresis-based microfluidic profiling of leaf protoplast subpopulations enables label-free differentiation of closely related herbal species, achieving high-confidence discrimination between moso and henon bamboo and demonstrating the utility of biophysical single-cell signatures for authentication [65].

The regeneration of the primary cell wall, a complex developmental process, has also been investigated using microfluidic impedance-based approaches. Chen et al. designed a platform combining microstructured electrodes with microfluidic channels for non-invasive, label-free quantitative assessment of PCW regeneration in single cells. Their system successfully discriminated between wild-type Arabidopsis cells and mutants with altered PCW properties (impairing the synthesis of cellulose and causing structural deregulation in PCW), highlighting the device’s potential for physical phenotyping [60].

A distinct application of microfluidics was presented by Zhang et al., who developed a plant-wearable microfluidic sensor for in situ detection of methyl parathion, a toxic but widely used pesticide. The device integrates capillary-driven electrolyte flow through microchannels and laser-induced graphene–gold nanoparticle electrodes. Attached directly to plant leaves, the sensor achieved high sensitivity across a wide concentration range (0.001 µM to 200 µM), offering rapid, continuous environmental monitoring capabilities [61].

Following the development of microfluidic analytical devices for biochemical and physiological assessments at the single-cell level, a complementary set of microfluidic platforms has emerged, specifically designed to investigate the biomechanical properties of plant cells. Understanding the mechanical behavior of plant cells is crucial, as these properties underpin cell function, environmental adaptation, and development, and can inspire biomimetic designs in material science and engineering.

Distinct from the previously discussed analytical tools, microfluidic devices for studying cell biomechanics often employ specialized structures that enable controlled application of mechanical forces to single cells. Plant cells exhibit a wide range of biomechanical properties, which reflect their functional specialization within the organism. Tip-growing cells, such as pollen tubes and root hairs, are particularly well-suited for these investigations due to their naturally invasive growth patterns and the mechanical challenges they encounter in vivo. Defining and quantifying these properties provides critical insights into cellular adaptation, developmental processes, and potential limitations imposed by environmental mechanical constraints. Analytical tools and microstructures used in cell biomechanics investigations are summarized in

Table 5. Microfluidic systems and structures enabling description of plant cell biomechanics.

Microdevice	Investigated Tissue, Organism	Function	Structure Characteristics	Measured Parameter	Ref.
Bending-Lab-On-Chip (BLOC)	Pollen tube, Camellia japonica	Bending of the pollen tube through fluid loading	Twisted growth microchannel to prevent backward movement of pollen, control channel and bending test chamber for measurement	Bending and rotational deflection Determination of the Young’s modulus	[66]
Flexure integrated Lab-on-a-Chip (FiLoC)	Pollen tube, Camellia japonica	Guiding pollen tube against microcantilever	Pollen grain is trapped and the pollen tube is guided through a growth channel against a microcantilever for invasive growth force measurement	Growth force, Growth dynamics upon interaction with a mechanical obstacle	[67]
Lab-On-Chip device	Pollen tube, Lilium longiflorum	Indentation of pollen tubes	Trapping microvalve for pollen grain immobilization, pollen tube growth channel with indentation microvalve	Compression and stretch ratio characterization	[68]

and illustrated in Figure 5.

The examples summarized in Table 5 share a common model system: pollen tubes growing inside microfluidic channels. Pollen grains are introduced into a main cultivation chamber, where they are immobilized using hydrodynamic traps and maintained by a continuous fluid flow. Channel geometries are often optimized to ensure secure immobilization of the pollen grains. Following germination, the pollen tubes grow through microchannels slightly larger than their diameter, facilitating mechanical interrogation. Depending on the experimental design, different mechanical forces are applied: compression or bending stresses are introduced via integrated test chambers [66,68], while microcantilevers serve as mechanical obstacles that allow for real-time measurement of growth forces and dynamic behavior [67]. These systems enable the quantification of critical biomechanical parameters such as Young’s modulus, invasive growth force, turgor pressure, and wall stiffness at the single-cell level.

The microfluidic systems summarized in Table 2, Table 3, Table 4 and Table 5 illustrate distinct trade-offs between environmental control, throughput, and long-term culture stability in plant cell–based investigations. Chamber- and microstructure-based platforms provide highly stable microenvironments that are well suited for extended cultivation, live-cell imaging, and biomechanical studies, but typically offer limited throughput and flexibility once fabricated. In contrast, microchannel-based devices enable precise spatial guidance and dynamic stimulus delivery, though prolonged confinement and shear stress may restrict long-term cell viability for sensitive plant cell types. Droplet-based and analytical microsystems excel in high-throughput screening and single-cell characterization, enabling rapid sampling and parallelization; however, maintaining droplet stability and sustained physiological relevance over extended timescales remains challenging. Together, these platforms are best viewed as complementary tools, with platform choice dictated by the balance between experimental duration, mechanical sensitivity, analytical depth, and throughput requirements.

3.2. Microfluidic Devices for Plant Development and Structural Characterization Research

Microfluidic devices offer platforms for live imaging, metabolite sampling, environmental modulation, and high-throughput phenotyping, often surpassing the limitations of traditional agar-based or soil-based methods. They have been particularly impactful for model plants like Arabidopsis thaliana, allowing for detailed investigations at both cellular and whole-plant levels. At the whole-plant scale, microfluidics can support root-zone management, spatially controlled chemical delivery, and real-time phenotyping under dynamic environmental conditions, facilitating integrated studies of growth, development, and stress responses. We categorized the devices developed for studying plant development and structural characterization based on their primary application areas: (1) microscale platforms for cellular and subcellular developmental studies, (2) high-throughput phenotyping, (3) root development and morphodynamics, (4) chemical and metabolite profiling, and (5) integrated environmental simulation platforms. An overview of selected studies is summarized in

Table 6. Microfluidic devices for plant morphogenesis and phenotyping studies.

Phenotyping Target	Device Design	Functionality	Measured Parameters	Plant Species	Refs.
Tissues growth and development	Growth chamber (30 µm deep and volume of 1.36 µL) with central inlet sector and surrounding flow control channels, bonded to a coverslip	Long-term live imaging	Growth rate, Cell expansion rate—area, length, width, cell division, Cytoskeleton degradation	Physcomitrella patens	[69]
Root and shoot growth and development	Chip with seed holding sites, root and shoot growing regions and 1.8 mm channel for media flow	Live imaging and monitoring of root and shoot phenotypes	Root length, hypocotyl length, cotyledon surface area	Arabidopsis thaliana	[70]
Root and shoot growth and development (up to 30 days)	Miniature greenhouse—plant chip with light intensity and temperature sensor in insulated space	Environmental control	Root length, hypocotyl length,	Arabidopsis thaliana	[71]
Germination and shoot growth	Plant array chip—300 2 × 2 mm square arrays grouped in 5 regions	Monitoring multiple replicants phenotypes, nutrient gradient control	Germination rate, radicle length	Arabidopsis thaliana	[72]
Germination, growth and etiolation	Plant array chip with 384 wells	Light gradient control	Germination rate	Arabidopsis thaliana	[73]
Root morphology	Microchannel-based device with channels width 150–400 µm	Growth space definition, live imaging	Root length, root diameter root hair length, number of root hair	Arabidopsis thaliana	[74]
Root hair nucleus morphodynamics	1.5 cm-long channels for main root growth (250 µm wide and 100 µm deep) connected via two arrays of lateral 400 µm-long perpendicular channels for root hairs expansion (20 µm wide and deep)	Live imaging	Position of the tip, tip average speed, nucleus to tip position	Arabidopsis thaliana	[75]
Spatial and temporal profiling of root exudates	Main channel for root growth, nanoporous membrane and two channels for sampling fluidics	Live imaging, Metabolite sampling	Metabolite concentration, diffusion	Arabidopsis thaliana	[76]
Gene expression dynamics	RootArray—growth chamber with 64 wells, liquid and gaseous chambers	Automated live imaging, environmental control	Fluorescence intensity (indicating gene expression)	Arabidopsis thaliana	[77]
Root growth and development	Root-TRAPR system—internal oval root growth chamber with transparent walls for microscopic observation and an external structural frame	Live imaging	root length, root surface area, average root diameter	Cannabis sativa	[78]
Root growth and development	RootChip—eight individual microchannels for root growth and observation 800 µm wide and 100 µm high	Live imaging, Förster Resonance Energy Transfer (FRET) measurements	Intracellular sugars (Glc, Gal) levels, root length, growth rate	Arabidopsis thaliana	[79,80]
Root exudation dynamics in soil-like environments	Rhizosphere-on-a-chip with synthetic porous structure, coupled with liquid micro-junction surface sampling probe mass spectrometry (LMJ-SSP-MS)	Long-term growth and imaging of roots, visualization and chemical analysis of root exudate distribution via mass spectrometry	Spatial distribution of amino acid hotspots	Brachypodium distachyon	[84]
Metabolomic profiling	microfluidic capillary zone electrophoresis-mass spectrometry (CZE-MS)	High-throughput alkaloid analysis with minimal sample prep; targeted and untargeted MS/MS	Relative abundance of alkaloids, electropherogram features, mass spectra	Lobelia cardinalis	[85]

.

Microscale microfluidic platforms have enabled detailed observation of developmental processes that are otherwise inaccessible in conventional culture systems. A notable example is the PDMS chamber system developed by Bascom et al., which supports long-term cultivation and high-resolution live imaging of Physcomitrella patens tissues. Because the chambers are directly bonded to a coverslip, the device allows continuous visualization of protonemal growth, subcellular dynamics, and mutant phenotypes over extended periods, with growth rates comparable to those on solid medium. This work demonstrates how stable microscale confinement paired with optical accessibility can facilitate real-time analysis of cellular and subcellular developmental events [69]. Building on such microscale approaches, a second class of devices—microfluidic chip arrays—focuses on high-throughput phenotyping of seeds and seedlings under controlled environments.

Simple yet highly useful microfluidic devices, such as chip arrays, have been developed to facilitate plant phenotyping by enabling faster workflows and easier comparison between experimental groups. These devices are primarily used for seed germination assays across different genotypes or culture conditions, seedling cultivation, and evaluation of root and/or leaf development. Early microdevices include chips with hydrodynamic seed traps [70] and multi-well arrays accommodating up to nearly 400 plantlets, where seeds are placed using an array seeder [72] or micro-vacuum seeder [73]. The ability to load large numbers of seeds, particularly small ones like Arabidopsis thaliana (250–600 µm in diameter), significantly reduces manual labor and minimizes mechanical damage.

Jiang et al. developed a device supporting 26 plants across two connected layers, where seeds are positioned in funnel structures and exposed to a constant flow of liquid medium through a 1.8 mm diameter channel. The relatively long growth channels enabled root imaging for up to 22 days; however, all seedlings were subjected to uniform growth conditions as they shared the same media flow [70]. This platform was later advanced by integrating it into a miniature greenhouse equipped with a Light-Emitting Diode (LED) ring, mini-fan, temperature and light sensors, and a control circuit allowing for rapid adjustments to the growth environment. A polymer-dispersed liquid crystal film on the front window enabled external imaging without disturbing the internal conditions [71].

Park et al. proposed a more complex and high-throughput system: a 20 × 20 array of 2 × 2 mm cultivation chambers. Seeds were distributed into agar-filled chambers using an array seeder, and the chip accommodated up to five different media compositions across the array. Seedling growth was monitored via digital single-lens reflex (DSLR) imaging [72]. Later, the system was further refined to be compatible with a standard 384-well plate, with seed loading optimized using a home-made micro-vacuum seeder featuring 16 suction needles. This modified array was used to study the effects of white light intensity on Arabidopsis growth [73].

These cultivation arrays exemplify the growing class of minifluidic and medifluidic devices—platforms whose dimensions are tailored to the physical characteristics of larger plant structures such as seeds, roots, or entire seedlings. While they depart from traditional microfluidic dimensions, these systems still rely on microfabrication principles and fluid control to achieve high-throughput phenotyping, parallel treatment conditions, and imaging compatibility. As such, they bridge the gap between chip-based microfluidics and whole-plant assays, providing a scalable approach to plant phenotyping.

Roots, due to their opacity and subterranean growth, are challenging research targets, yet their behavior provides critical insights into whole-plant health. Several microfluidic platforms have been developed for continuous live observation and detailed morphodynamic studies of plant roots.

Grossmann et al. introduced RootChip, a microchannel-based device enabling simultaneous cultivation and imaging of up to eight Arabidopsis plants. The device’s geometry was optimized for main root growth, and key parameters such as root length, growth rate, and intracellular sugar levels were monitored [79]. Another high-throughput device, RootArray, supported the growth of 64 plants in a configuration with roots in liquid media and shoots exposed to gaseous conditions. Integration with computational tools allowed for live imaging and automated analysis of gene expression patterns in 12 different transgenic lines under varying conditions [77].

Focusing on microchannel geometry, Sun et al. developed a crossed-channel device optimized for root diameter and growth length, which was used to investigate the effects of high salinity and phytohormone treatments on root and root hair development [74]. Singh et al. presented a more sophisticated coverslip-based chip, composed of three individual devices each supporting one plant. Each device included a main channel for primary root growth and numerous narrow side channels designed to individually guide root hairs, enabling precise tracking of nuclear migration during root hair development [75].

Alternatively, Suwanchaikasem et al. introduced Root-TRAPR, a modular growth chamber featuring an oval design that allows roots to develop more naturally, avoiding the restrictive geometry of typical microchannels. This system supported studies on the effects of chitosan treatment on root development, combining live imaging with root exudate collection, metabolic profiling, and genetic analysis [78].

Functional phenotyping, which encompasses chemical and metabolite profiling, remains a relatively underexplored aspect of plant analysis. These studies offer insight into dynamic biochemical changes in plant tissues in response to environmental conditions.

Patabadige et al. developed a system wherein plant roots grew within microchannels, while exudates were sampled through a perpendicular network separated by a nanoporous membrane. This allowed continuous, non-invasive collection of root exudates for spatiotemporal analysis via mass spectrometry, enabling the identification of compounds such as alanine, 5-oxoproline, isoleucine, and sucrose [76].

Aufrecht et al. introduced the rhizosphere-on-a-chip, a device mimicking natural soil structure with synthetic sand grains of varying sizes, allowing roots to explore a semi-natural microenvironment while retaining optical transparency for live imaging. This platform facilitated the prediction and experimental validation of root exudate hotspots using liquid microjunction surface sampling probe mass spectrometry (LMJ-SSP-MS) [84].

Alternatively, microfluidic technologies can be employed solely for metabolite analysis rather than cultivation. For example, microfluidic capillary zone electrophoresis coupled with mass spectrometry was used to analyze alkaloid levels in Lobelia cardinalis hairy root extracts, demonstrating another application of microscale techniques in plant metabolic studies [85].

Microfluidic devices developed for plant morphogenesis and phenotyping (Table 6) offer improved control over growth orientation, nutrient supply, and imaging conditions compared to conventional cultivation systems. Platforms based on structured chambers or guided growth channels support reproducible phenotyping and real-time monitoring of developmental dynamics, particularly for roots and seedlings. However, scalability toward mature plants remains limited by chamber size, nutrient delivery constraints, and the difficulty of integrating aerial tissues within enclosed microenvironments. While larger mesofluidic and modular designs partially address these challenges, trade-offs between environmental control and biological realism persist. Consequently, such platforms are most effective for early developmental stages and comparative phenotyping rather than full life-cycle studies.

3.3. Devices for Investigating Plant Cell Stress Response and Communication

The integration of microfluidic systems into plant biology research has transformed our ability to study plant responses to abiotic stresses with unprecedented precision. These devices allow for the manipulation of environmental variables, such as water availability, nutrient gradients, salinity, toxic additives and mechanical forces, while enabling continuous imaging and molecular assays. Particularly, root tissues have been extensively investigated using microfluidic platforms because they are the primary sites of environmental perception and signaling.

Table 7. Microfluidic devices used in investigation of signaling during abiotic stress in plants.

Type of the Stress	Investigated Tissue, Organism	Cultivation Condition	Microdevice	Analysis Method	Refs.
Drought	Roots, Arabidopsis thaliana	Media supplemented with 10 or 20% PEG-6000	RootChip	Light microscopy	[86]
Salinity, Deficiency of Pi	Roots, Arabidopsis thaliana	Media with 100 mM NaCl, Media with deficient Pi (0.01 mM)	Dual-Flow-RootChip	Calcium imaging—fluorescence Microscopy Light microscopy	[87]
Salinity, Drought	Roots, Arabidopsis thaliana	100 mM NaCl, 20% PEG-6000	Bi-directional-dual-flow-RootChip	Calcium imaging—fluorescence microscopy	[88]
Nutrient flow	Primary root, Brassica juncea	Nutrient flow ranging from 0 to 1.2 mL/hr	Microfluidic channel	Light microscopy	[90]
Drought	Roots, Oryza sativa	0,6% agar culture media containing 0%, 5%, 10%, 20% or 30% PEG-6000	Multi-chamber petaloid root-growth chip	Light microscopy, Gene expression analysis	[95]
Salinity, Drought	Roots, Nicotiana tabacum	150 mM NaCl, 10% PEG-6000	Foldable Plant Array Chip	Light microscopy	[97]
Microgravity	Seed, Arabidopsis thaliana	Treatment of seeds with IAA, Kyn and Gd3+	Microfluidic chip with 5 channels for seed cultivation	Light and fluorescence microscopy	[98]
Drought	Roots, Brachypodium distachyon	20% PEG	Microfluidic channel	Light microscopy	[89]
Deficiency of Pi	Leaf extracts	Media with deficient Pi (0.05 mM)	Multichannel microfluidic chip with a detection area	Sandwich hybridization—fluorescence detection of miR399	[91,92]
Drought	Plant extract, Arabidopsis thaliana	Plants cultivated in pots with dry soil	Paper-based microfluidic sensor	Colorimetric proline detection	[100,101]
Drought	Leaf stomata, Spathiphyllum wallisii	Plants cultivated in pots with soil with or without watering	Electro-mechanical sensor	Optical stomatal aperture Measurement Electrical and Raman measurements	[102]
Heat	Seed and root, Arabidopsis thaliana	Seeds placed in water, temperature in the range 25.3 °C—37.2 °C	Microfluidic Seed Growth Chips (SGC) incorporated with electrospun nanofibrous membranes (ENMs)	Light microscopy	[93]

provides an overview of the current microfluidic devices applied in the study of plant signaling under abiotic stress, highlighting their cultivation methods, stress types, analytical approaches, and major findings. Together, these platforms offer versatile, scalable tools for advancing our understanding of plant stress biology.

Many studies on the abiotic stress response in plants have led to the establishment or improvement of microfluidic devices. As described in the previous chapter, the RootChip platform has been further developed and modified to allow specific environmental control and analysis. Stanley et al. introduced a dual-flow version of this device, enabling root growth in a laminar flow environment of two fluid compositions. This setup allows real-time observation of the effects of stress exposure to only one side of the root. Additionally, the main growth channel was equipped with a triangular pillar array to ensure straight root growth and even exposure to each medium. Using this device, the effects of asymmetric phosphate exposure (Pi), as well as drought stress (induced by 20% PEG 8000), on root hair growth were investigated [87]. Further improvements were presented by Allan et al., who added a bi-directional flow feature. This modification enabled not only control over the fluid composition on either side of the Arabidopsis primary root but also directional flow control, allowing selective exposure of the root tip to a desired stress factor. The platform’s usability was demonstrated through studies on calcium signaling in response to salinity (NaCl-induced) and drought (PEG-induced) stress [88].

Moussus and Meier modified the original TipChip device by demonstrating the potential of 3D printing as a fabrication method. They tested the biocompatibility of various 3D printing materials with Arabidopsis thaliana and ultimately selected GR-10, despite observing a decreased germination rate for all tested materials. The final device could accommodate the growth of six individual plants and was used to investigate drought stress (induced by 10% and 20% PEG-6000) through analysis of root growth and development [86].

Several other devices, similar to the TipChip platform, have been developed over the years. These devices typically consist of a single microfluidic channel supporting primary root growth under a continuous fluid flow with variable effectors. Khan et al. created a simple device comprising six separate channels for root growth. Using fluorescence microscopy, they visualized the maturation zone, root tip, and cross-sectional zones of Brachypodium distachyon roots under 24 h osmotic stress (20% PEG) conditions. They also performed gene expression analysis of several up- and downregulated genes associated with osmotic pressure [89]. Similarly, Agarwal et al. proposed a microfluidic setup for supporting Brassica juncea root growth with a constant nutrient flow through the microchannel. They determined root growth rate under various medium flow rates and studied nutrient uptake under flow-modulated mechanical stress [90].

Another approach to stress investigation is a foldable microfluidic chip array used for salinity (NaCl-induced) and drought (10% PEG) studies. This device consists of multiple chips, each containing several seed germination chambers and corresponding conical-shaped root growth chambers. The chips are connected by elastic hinges, allowing for rapid relative comparison of plant phenotypes, such as germination rate and root length [97].

Applying a range of effector concentrations to different parts of the same organ provides highly reliable results by limiting experimental variation to the tested substance alone. Chai et al. demonstrated a microfluidic petaloid root-growth chip consisting of five separate growth chambers, each supporting a root portion from the same Oryza sativa plant. In their study, drought stress was applied using 5%, 10%, 20%, and 30% PEG-6000, and root length, shoot length, and total root projection area were determined via optical imaging [95].

A distinct study focusing on microgravity-induced stress was conducted by Du et al. They used a microfluidic chip containing five cultivation chambers for Arabidopsis thaliana seeds, with permanent magnets placed underneath. Seeds were subjected to microgravity conditions via Gd³⁺-promoted magnetic levitation, and germination and auxin distribution were assessed using fluorescence microscopy [98].

Beyond controlled lab conditions, microfluidic technologies have been adapted for use in natural environments to monitor plant stress directly. A microfluidic device for fluorescence detection of miRNAs was proposed for the early detection of phosphorus deficiency in leaf extracts. Based on a sandwich hybridization method, this platform enables rapid and sensitive detection (limit of 1 nM) of the Pi-stress-related miR399c biomarker [91,92]. Choi et al. introduced a paper-based analytical device for diagnosing drought stress through colorimetric detection of proline. This sensor, fabricated from Whatman paper patterned with wax and loaded with ninhydrin, reacts with proline present in plant extracts [100]. Further improvement was achieved by adding a second paper layer, enhancing field applicability with a measurement range of 3–50 mM and a detection time of only 3 min [101].

Another type of sensor for persistent drought monitoring was developed by Koman et al. They presented an electro-mechanical sensor based on a microfluidic chip attached to a plant leaf to monitor stomatal behavior. Stomata are microscopic pores on the leaf surface that regulate gas exchange and water loss by opening and closing in response to environmental cues such as light, humidity, and soil moisture. By applying a nanoparticle-based conducting ink, the sensor could differentiate between open and closed stomata states, correlating to illumination and soil water status [102].

Lastly, a microfluidic seed growth chip coupled with an electrospun nanofibrous membrane (ENM) was developed for evaluating heat stress responses. This device consists of 20 seed chambers arranged in a 4 × 5 array and interconnected by microchannels. Beneath the microfluidic layer, an ENM layer allows precise temperature control. Root development under heat stress was assessed through measurements of root length and width [93].

While microfluidic platforms for studying abiotic stress have provided valuable insight into plant responses at the level of individual cells, tissues, and organs, understanding plant biology requires an equally detailed exploration of how cells communicate both within the plant and with external organisms. In this section, we highlight microfluidic devices specifically designed to investigate cell–cell communication, offering new perspectives on intra- and interspecies interactions under controlled conditions.

Most of the devices discussed so far were developed for the study of relatively uniform cell populations, where measured parameters were typically averaged across large groups or restricted to limited subpopulations. In contrast, the systems described here are designed to accommodate different types of cells and focus on the interactions between them. Some devices provide insight into communication within a single organism, while others investigate interactions with symbiotic or pathogenic microbes. These devices are summarized in

Table 8. Characterization of microfluidic devices for cell–cell communication.

Device	Type of Cell–Cell Interaction	Device Structure	Experimental Multiplicity	Investigation Method	Refs.
Modular microfluidic bioreactor	Somatic cell–cell (cell line BY-2, N. tabacum)	Connectable chips containing 800 µL cell chamber and perfusion chamber separated by nanoporous membrane	1, with possibility of multiple cell types combined in separate chambers	Proliferation factor measurement	[103]
	Metabolic synergy between somatic cells (cell strains from seedlings of Catharanthus roseus)			HPLC–DAD-ESI–MS/MS for alkaloid detection
	Fungal phytotoxicity (cell line BY-2, N. tabacum an Neofusicoccum parvum)			Cell mortality assay
Microfluidic trapping device	Plasmodesmata permeability between tobacco BY-2 cells	Channels for trapping plant filaments	1 cell population	Confocal microscopy, Fluorescence Recovery After Photobleaching (FRAP) measurement	[104]
Tracking root interaction system (TRIS)	Root-bacteria (Arabidopsis thaliana and Bacillus subtilis)	9 channels of 160 µm height, each with 3 individual ports: inlet and outlet for bacteria introduction and one for introduction of germinated seeds	9	Light and fluorescence microscopy	[105]
Channel-based device	Root-bacteria colonization (Arabidopsis thaliana and strains: Pantoea sp. YR343 and Variovarax sp. CF313)	Main channel for root growth 150 µm high, 200 µm wide and 3.8 mm long, surrounding treatment chamber 20 µm high and 8 areas with injection channels	1	Light and fluorescence microscopy	[106]
Imaging EcoFab	Root-microbes interactions (Brachypodium distachyon and Pseudomonas simiae strains)	Oval chamber covered with pillars, allowing for media flow while flat root growth against the coverslip	1	confocal microscopy	[107,108]
Y-channel device, Open format and Soil-analog microfluidic device	Root-bacterial biofilm (Populus trichocarpa and Pantoea YR343)	Devices allowing for the growth of the roots and formation of rhizosphere, with live sampling of root exudates	1	Mass spectrometry	[109]
Plant-in-chip microchannel based device	Root-pathogens (Arabidopsis thaliana interacting with sugarbeet nematode or Phytophtora sojae)	8 parallel straight microchannels 80 µm high, 350 µm wide, 1 cm long, connected with thin vertical channels for pathogens introduction	8	Light microscopy	[110]
RMI-chip	Root-microbe interactions (roots of Populus tremuloides and Pseudomonas fluorescens)	Root growth channels 100 µm high, 800 µm wide and 36 mm long with two inlets for media and bacteria inoculation	12	Light and fluorescence microscopy	[111]
Channel-based device for root growth	Root-pathogens (Arabidopsis thaliana and Phytophthora parasitica) kinetics of zoospores in the vicinity of the root	Root growth channels 150 µm high and of various width: 0.5, 1, 2 or 4 mm	up to 8	Light microscopy	[112]
Miniaturized plant-microbial fuel cell (mPFMC)	Root exudates as carbon source for electricity generating bacteria (rice plants and bacterial strains: Shewanella oneidensis MR-1 and Pseudomonas aeruginosa PA14)	Chamber for hydroponic plant growth, carbon cloth for bacteria growth, separated from each other via semipermeable filtering membrane	1	Electrochemical measurements, GC-MS for root exudates analysis, SEM for bacterial biofilm observation	[116]
Droplet-based single nucleus RNA sequencing (sNucRNA-seq) platform	Root-symbiotic bacteria (Glycine max and Bradyrhizobium diazoefficiens USDA 110)	Microfluidic channels included in the sequencing platform	High-throughput analytical platform	RNA sequencing	[117]

.

A critical design consideration in microfluidic devices used for investigating cell–cell interactions is the degree of contact and interference between interacting organisms. This is primarily determined by the spatial separation of cultivation chambers. Depending on the intended focus of the study, interacting organisms can either be separated by a semi-permeable membrane—facilitating metabolic exchange without direct contact—or co-cultivated in a shared chamber to allow detailed observation of physical interactions and signaling mechanisms. Chemical signaling involves the diffusion of metabolites, hormones, or other molecular signals across media to influence neighboring cells, while physical or mechanical communication requires direct contact, such as cell adhesion, biofilm formation, or structural interactions like hyphal penetration or root colonization.

In plant–microbe interaction studies, direct interactions are often explored at the root–microorganism interface, with roots typically cultivated in microchannels or chambers into which microorganisms are introduced and allowed to move freely. One of the first microfluidic devices for this purpose was described by Parashar and Pandey. They developed a system comprising a grid of eight separate channels, each supporting the growth of a single Arabidopsis thaliana seedling, thereby enabling parallel investigation of eight replicates while minimizing cross-contamination. Each microchannel was connected to thinner vertical channels for the introduction of chemicals or pathogens. Using this setup, the authors demonstrated early stages of pathogen–host interaction by inoculating zoospores of Phytophthora sojae and sugar beet nematodes (SBNs) into the channels and monitoring their effects under light microscopy over four days [110].

A comparable device was introduced by Massalha et al., termed the Tracking Root Interaction System (TRIS). This system accommodated nine separate roots, each growing in an individual channel mounted directly onto a microscopic stage. The channels were designed at an angle, promoting vertical growth of the seedlings before allowing roots to transition into a horizontal plane for improved imaging. Fluorescently labeled strains of Bacillus subtilis were introduced through inlets at the channel ends, and real-time colonization of root surfaces was observed using dark-field microscopy over a 30 min period. Longer observations, up to 12 h, were conducted using confocal microscopy, revealing bacterial accumulation at the root tips. Moreover, the system enabled the study of bacterial competition by co-inoculating roots with GFP-labeled Escherichia coli and mKate-labeled Bacillus subtilis strains [105].

Building on these designs, Noirot-Gros et al. adapted a similar multi-channel device to accommodate larger seedlings of Populus tremuloides. Their platform featured individual ports for inoculation, allowing the introduction of Pseudomonas fluorescens strains that formed biofilms on the root surfaces. Continuous imaging by laser-scanning confocal microscopy revealed diverse bacterial assemblies along the root surface [111].

A slightly different approach was taken by Aufrecht et al., who developed a device designed to investigate root–bacteria interactions at multiple root locations within a single plant. Although only one Arabidopsis thaliana seedling could be accommodated at a time, the device featured eight injection channels for targeted bacterial introduction along different regions of the root. After an initial cultivation period of three days, various fluorescently tagged bacterial strains were inoculated, and colonization was monitored over four days through fluorescence microscopy at 24 h intervals. This study revealed that the extent and pattern of root colonization were influenced by the initial bacterial inoculum density [106].

While microchannels are effective for structured root cultivation, alternative designs aim to better mimic the rhizosphere environment. Jabusch et al. developed the Imaging Fabricated Ecosystem (Imaging EcoFAB), an oval, transparent cultivation chamber capable of accommodating the entire root system of Brachypodium distachyon seedlings. To enhance imaging resolution and bring roots closer to the coverslip, the chamber floor was embedded with pillar structures, reducing the effective chamber height to approximately 1 mm. Roots growing freely between the pillars were visualized using light microscopy, and rhizosphere colonization was demonstrated using nine different strains of Pseudomonas simiae, each expressing a unique fluorescent protein. Additionally, the system was employed to study cross-species interactions, such as the growth of Neurospora crassa fungal hyphae along the roots [107,108].

Further exploration of plant–pathogen interactions was conducted by Cohen et al., who designed a microfluidic device with a single channel supporting primary root growth. In their study, droplets containing Phytophthora parasitica zoospores were introduced at one end of the channel, and zoospore movement trajectories were recorded at multiple locations along the root. They observed that zoospore speed and trajectory were significantly influenced by the presence of roots but only within a proximity of 300 µm [112].

The devices described thus far emphasize co-cultivation, providing valuable insights into symbiotic colonization and pathogenic invasion. However, in studies focusing on indirect interactions—such as signaling via secreted metabolites—physical separation of organisms may be preferred. For this purpose, modular microfluidic bioreactors have been developed. One such system, described by Finkbeiner et al., comprises multiple modules connected by tubing. Each module contains two chambers: an upper cultivation chamber for cells and a lower perfusion chamber for media flow, separated by a semi-permeable membrane supported by a mesh. This configuration allows chemical exchange without cell migration between chambers. By arranging the modules sequentially, researchers could investigate how metabolites produced by one cell type influence another. Finkbeiner et al. demonstrated the system’s versatility across three examples: enhancing vincristine synthesis through metabolic synergy between two Catharanthus roseus strains; regulating proliferation in tobacco BY-2 cells via quorum sensing; and detecting phytotoxins produced by Neofusicoccum parvum, leading to tobacco cell death [103].

A similar principle underlies the miniaturized bioelectricity generation device presented by Jiang et al., which cultivated hydroponic rice plants alongside electrogenic bacteria (Shewanella oneidensis MR-1 and Pseudomonas aeruginosa PA14). In this device, root exudates served as a carbon source for bacterial growth on carbon cloth electrodes. A semi-permeable membrane separated plant roots from the bacteria, enabling indirect interaction and resulting in bioelectricity generation without direct physical contact [116].

Interactions also occur at even finer scales, between neighboring plant cells. Investigating these processes requires single-cell resolution. Kurotani et al. developed a microfluidic device for trapping tobacco BY-2 cell filaments expressing GFP to study plasmodesmatal permeability using fluorescence recovery after photobleaching (FRAP) [104].

Finally, plant–microbe interactions not only manifest at cellular and structural levels but also induce genetic and transcriptomic changes. D’Agostino et al. utilized root nodules and tissues from Glycine max to perform single-nucleus RNA sequencing (sNucRNA-seq) using a droplet-based microfluidic platform. This technique enabled high-resolution dissection of molecular mechanisms underpinning symbiotic interactions [117].

In addition to direct interactions between different organisms or populations, communication within plant tissues themselves is often mediated by chemical gradients. One prominent example is chemotropism—the directional growth of plant organs in response to external chemical signals. Chemotropism plays a crucial role in diverse aspects of plant biology, from root foraging for nutrients to the guidance of pollen tubes during fertilization.

Microfluidic systems offer unparalleled precision in controlling the spatiotemporal distribution of chemical cues, enabling the generation of stable gradients that closely mimic natural conditions. These platforms facilitate real-time observation of plant responses, providing valuable insights into the signaling pathways, molecular mechanisms, and cellular behaviors underlying chemotropic phenomena.

A particularly well-studied case is pollen tube guidance, where microfluidic devices featuring arrays of channels and chambers are employed to trap ovules while positioning pollen tubes at a central point. This setup allows researchers to monitor pollen tube growth direction and attraction dynamics in response to chemical signals emitted by the ovules. Such devices are summarized in

Table 9. Microfluidic devices for chemotropic plant cells activity investigation.

Type of the Device	Purpose	Characteristics	Cell Type, Organism	Ref.
Microsystem-based microfluidic device simulating ovule microenvironment	Assessment of pollen tube guidance in response to ovule-derived cues	Main groove (1 mm wide, 5 mm long) with side chambers (250–1000 µm2) for ovule placement or gradient generation, depth along whole device around 500 µm	Pollen tubes, Arabidopsis thaliana	[113]
T-junction and crossroad microchannel devices	Assessment of pollen tube guidance in response to ovule-derived cues	Microchannels of 500 µm in width and 25 µm in height. The distance between the center of style inlet and: T-junction—3.5 mm; crossroad—2 mm	Pollen tubes, Torenia fournieri	[114]
Microslit-based microfluidic chip	Quantitative assessment of pollen tube chemotropism	Thin microslit channel array (2–16 µm in width and 5 µm in height) and thick channels (90 µm in height) with style inlet, sample reservoir (containing ovary) and blank reservoir	Pollen tubes, Torenia fournieri	[115]

.

Yetisen et al. developed a microsystem-based assay that closely mimics the in vivo microenvironment of ovule fertilization in Arabidopsis thaliana. This platform allowed quantitative assessment of pollen tube growth, targeting frequency, and directional preference toward unfertilized ovules, confirming the ability of pollen tubes to sense and respond to minute chemoattractant gradients [113]. Sato et al. expanded this approach by implementing T-junction and crossroad microchannel devices for Torenia fournieri pollen tubes. These devices provided a controlled pathway for pollen tube growth, achieving guidance response ratios of 56–57% and enabling the collection of pollen tubes attracted to female tissue with high purity for downstream molecular analyses [114]. Further refinement was achieved by Yanagisawa and Higashiyama, who introduced microslit-based channels to selectively permit only chemoattractant-responsive pollen tubes to reach the ovule source. This approach allowed precise quantification of chemotropic responses, excluding randomly growing pollen tubes and highlighting the potential of microfluidic selection for functional studies on signaling mechanisms [115]. These studies collectively demonstrate the versatility and precision of microfluidic devices in dissecting chemotropic responses in plant cells.

Microfluidic platforms developed for investigating abiotic stress responses and cell–cell communication in plants (Table 7, Table 8 and Table 9) predominantly focus on root systems and polarized tip-growing cells, such as pollen tubes, where precise spatial control of the microenvironment is essential. Root-based devices typically combine microchambers with guided microchannels to enable stable long-term cultivation while allowing localized application of chemical, mechanical, or osmotic stimuli. These configurations are well suited for monitoring adaptive stress responses, hormone signaling, and root–microbe interactions over extended timescales, although the confined geometry may limit the representation of complex soil heterogeneity. In contrast, microchannel-based platforms designed for pollen tube studies provide high temporal resolution and directional guidance for analyzing chemotropism and cell–cell signaling, but are generally restricted to short-term experiments and simplified extracellular environments. Overall, these systems prioritize environmental precision and imaging accessibility over throughput, making them particularly effective for mechanistic studies of plant stress signaling and intercellular communication.

4. Future Perspectives: Harnessing Microfluidics for Sustainable and Scalable Plant Technologies

Microfluidic systems are typically designed for specific research purposes, tailored to the biological characteristics of the cells, tissues, organs, or organisms under investigation. With recent technological advancements, numerous platforms have emerged that are well-established and capable of addressing complex scientific questions in plant research. A key to increasing the versatility of microfluidic devices lies in further customizing system parameters, such as the number, geometry, and dimensions of channels, cultivation volumes, fluid flow dynamics, and the inclusion of auxiliary microstructures, to meet the needs of diverse biological models.

Plant technologies span a wide range of applications, from agriculture and biotechnology to environmental monitoring and conservation. Each of these fields presents distinct challenges that demand innovative, precision-oriented tools. Microfluidics, grounded in the precise control of microscale fluid volumes, offers a compelling suite of capabilities to address these needs. However, many currently available systems are highly specialized and limited in scope, underscoring the necessity for continued innovation.

In sustainable agriculture, microfluidic devices have the potential to revolutionize water and nutrient delivery. Conventional irrigation techniques often result in overuse and environmental degradation, whereas microscale systems can optimize resource distribution at the root zone, thereby reducing waste and improving crop yield. Similarly, the impacts of climate change, including temperature shifts and altered precipitation patterns, call for resilient crop varieties. Microfluidic platforms can simulate dynamic environmental conditions to screen stress responses, accelerate trait selection, and support breeding efforts.

Microfluidics also presents valuable tools for combating plant diseases. By enabling the controlled introduction of pathogens and monitoring of immune responses, these systems allow for detailed characterization of plant–microbe interactions. This capability is crucial for developing next-generation disease control strategies. Additionally, in the context of soil health, microfluidic devices can be used to assess microbial communities, nutrient availability, and pollutant concentrations—insights that can be further leveraged to optimize phytoremediation processes.

In plant biotechnology, microfluidics facilitates gene editing, transformation, and tissue regeneration within tightly controlled microenvironments. These systems enable more efficient development of genetically modified crops with improved resistance to environmental stressors and pathogens. Beyond laboratory use, microfluidic sensors emerge as essential tools for environmental monitoring. Their ability to detect stress markers and contaminants in real time supports decision-making in land management and conservation strategies.

A particularly promising but technically challenging frontier lies in developing soil-on-a-chip systems capable of sampling exudates and nutrient fluxes without disturbing the microenvironment. This approach requires ultra-sensitive methods, such as microjunction-coupled mass spectrometry, to capture spatial and temporal data without disrupting native chemical gradients.

Despite these opportunities, several technical and biological barriers remain. Plant cells, with their rigid walls and complex architecture, behave differently from animal cells under fluidic stress. This necessitates the design of highly specialized platforms capable of mimicking the mechanical and chemical properties of plant tissues. Maintaining long-term cell viability within microfluidic systems is another major challenge, particularly for protoplasts and other fragile cell types that require finely tuned conditions.

Scalability continues to be a critical obstacle. While high-throughput microfluidic assays demonstrate strong potential under controlled laboratory conditions, their translation into robust, field-deployable agricultural technologies remains limited. Many existing platforms lack sufficient modularity and adaptability to accommodate different plant species, developmental stages, and experimental objectives. Future progress is therefore likely to rely less on increasing architectural complexity and more on intelligent data integration. In particular, coupling microfluidic platforms with automated imaging, machine learning, and artificial intelligence offers a powerful route toward real-time data interpretation, phenotype classification, and decision-making. Recent advances highlight the growing role of AI-assisted lab-on-chip systems in the early detection of plant diseases, where microfluidics combined with image analysis and digital diagnostics enables rapid, sensitive, and low-cost identification of pathogenic threats [126]. Such approaches may help bridge the gap between laboratory-scale innovation and scalable, practical solutions for crop monitoring and plant health management.

Material limitations further hinder the broad adoption of microfluidics in plant studies. While PDMS remains a popular choice, it poses challenges in terms of chemical compatibility and optical properties. Development of new materials that are biocompatible, chemically stable, and ideally biodegradable will be key to future breakthroughs.

Finally, meaningful progress in this space depends on bridging disciplinary divides. Effective integration of microfluidics into plant research requires collaboration among plant biologists, engineers, chemists, and data scientists—each contributing to the refinement of experimental systems and interpretive tools.

5. Conclusions

Microfluidics has established itself as a transformative tool for plant physiology and development research, offering unmatched precision, integration, and adaptability at the microscale. Yet to fully realize its potential, particularly in advancing sustainable agriculture and climate resilience, researchers must address technical constraints, enhance system versatility, and foster interdisciplinary collaboration. By tackling these challenges head-on, the next generation of microfluidic platforms could redefine how we cultivate, protect, and understand both natural plant ecosystems and human-impacted environments, including agricultural and other artificial systems.