Simplified Sample Preparation and Lateral Flow Immunoassay for the Detection of Plant Viruses

TL;DR

A simplified lateral flow immunoassay with an easy sample preparation method enables rapid and sensitive detection of plant viruses like cowpea chlorotic mottle virus in the field.

Contribution

The paper introduces a simplified dipstick LFA format and an on-site sampling method for plant virus detection without laboratory equipment.

Findings

The assay can detect CCMV at concentrations as low as 3.5 µg/L in 15 minutes.

The new sampling method combines grinding, extraction, filtration, and reconstitution using a manual punch and syringe.

The assay detected systemic CCMV infection in cowpea plants before visual symptoms appeared.

Abstract

Lateral flow immunoassays (LFAs) are widely used for on-site testing; however, their use for the rapid detection of plant viruses in the field is often limited by inconvenient sample preparation. Here, we present a new sampling method and a simplified dipstick LFA format for the detection and monitoring of cowpea chlorotic mottle virus (CCMV) as a model plant pathogen. The assay employs a monoclonal mouse antibody for capture and a poly-clonal rabbit antibody conjugated to 80 nm gold nanoparticles for detection. Conventional sample and conjugate pads are omitted, allowing the test strips to be dipped directly into wells containing plant extract and antibody–gold conjugate. No plastic casing was required, which could lead to a reduction in waste. It was shown that CCMV concentrations as low as 3.5 µg/L or 350 pg per sample could be reliably detected in 15 min. Specificity tests confirmed that other plant viruses, cowpea mosaic virus (CPMV) and tobacco mosaic virus (TMV), did not produce false-positive results. In addition, we describe a new method for on-site sampling using a manual punch and a syringe equipped with a frit. This step combines grinding the sample, extraction, filtration, and reconstitution and mixing of the antibody-gold conjugate, enabling the analysis of punched leaf disks without laboratory equipment. When applied to CCMV-infected cowpea plants, the assay revealed systemic infection before visual symptoms became apparent. This work demonstrates that simplified LFAs combined with innovative sampling techniques can provide sensitive, specific, and rapid diagnostics for crop monitoring and support early intervention strategies in agriculture.

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Full text

1. Introduction

Lateral flow assays (LFAs) are one of the most popular immunoassay formats and have received enormous attention worldwide during the coronavirus pandemic. LFAs and similar technologies have been used and adapted for many areas. Their versatility has enabled applications across numerous fields, as summarized in several recent comprehensive reviews [1,2,3,4,5,6,7,8,9,10]. Building on this substantial literature, we focus here on specific technological challenges that still arise in practical LFA use, including workflow limitations and performance constraints [11]. This article contains some suggestions for overcoming these issues. We focused on three main objectives: (1) simplifying sample preparation, (2) improving user-friendliness, and (3) reducing manufacturing complexity and cost.

As a representative application, we selected a well-known plant pathogen, the cowpea chlorotic mottle virus (CCMV) [12]. The assay is designed as a field test that does not require laboratory equipment, such as homogenizers, centrifuges, pipettes, and readers. It does not even require water or electricity. In its current version, the assay can be completed in under 20 min, including sampling. There are quite a few lateral flow assays for plant pathogens available [13,14,15]. Indeed, LFAs have been developed for many plant viruses (e.g., TMV, PVY, and CMV) [16]. However, the range of assays specifically targeting CCMV appears to be very limited. CCMV infects cowpeas (Vigna unguiculata) and related species, which are predominantly cultivated in African countries, such as Nigeria, Niger, Burkina Faso, Kenya, and Senegal. On other continents, some production is reported in Asia, Central America, and South America, particularly in Brazil. Global production amounted to more than 9 million tons annually [17]. Unfortunately, cowpeas are affected by many pests and diseases, including cowpea mosaic virus (CPMV) and CCMV. Since plant viruses cannot be controlled with pesticides, early detection and subsequent removal of infected plants are the most effective countermeasures. In many resource-limited countries, economic constraints make systematic monitoring difficult. Therefore, the cost and user-friendliness of the respective tests are two of the most important aspects. Our work aims to address these constraints by developing a test that is highly accessible and easy to operate, requiring only minimal user guidance to ensure reliable performance.

2. Materials and Methods

2.1. Buffers, Chemicals, and Other Reagents

Buffers: running buffer (1× PBS with 1% (v/v) Tween 20, pH 7.4); carbonate buffer (sodium hydrogen carbonate (46 mM), sodium carbonate (54 mM), pH 10.0); and sodium acetate buffer (sodium acetate (30 mM), acetic acid (20 mM), Na_2_EDTA (1 mM), pH 4.8).

Chemicals and other reagents: PBS (cat. no. A0965,9010, AppliChem, Darmstadt, Germany); Tween 20 (cat. no. 11Tween201, MP Biomedicals, CA, USA); sodium hydrogen carbonate (cat. no. 8630, Th. Geyer, Renningen, Germany); sodium carbonate (cat. no. AP141648.1211, AppliChem, Darmstadt, Germany); sodium acetate (cat. no. 8694, Th. Geyer, Renningen, Germany); acetic acid (cat. no. 2289, Th. Geyer, Renningen, Germany); EDTA disodium salt di-hydrate (cat. no. 131669.1209, AppliChem, Darmstadt, Germany); defoamer (neodisher S Entschäumer, cat. no. 430148, Dr. Weigert GmbH, Hamburg, Germany); sodium azide (cat. no. 8690, Th. Geyer, Renningen, Germany); skim milk powder (cat. no. A0830, AppliChem, Darmstadt, Germany); liquid fertilizer (cat. no. 49128, Mairol GmbH, Gerstetten, Germany); ProClin 300 (cat. no. 48912-U, Sigma Aldrich, St. Louis, MO, USA); silicon carbide powder 600 mesh (cat. no. A13561, Alfa Aesar, MA, USA); household blender (Standmixer VitaBoost cat. no. MMBH6P6BDE, Bosch, Gerlingen, Germany); non-binding 96-well microtiter plate (white, cat. no. 655904, Greiner, Kremsmünster, Austria); 2 mL polypropylene syringe vessel (cat. no. V020PE061; MultiSynTech GmbH, Witten, Germany); handheld 6 mm hole punch (Universal Product Code: 795973640185, QWORK, distributed by Amazon, WA, USA); nitrocellulose membrane (Vivid 90 LFNC, 25 mm × 300 mm; cat. no. VIV902503R, Cytiva, MA, USA); LFA backing card (cat. no. 10547158, Cytiva Life Sciences, MA, USA); LFA absorbent pad (cat. no. 8115-2250, Cytiva Life Sciences, MA, USA); and gold nanoparticles for passive adsorption of polyclonal detection antibodies (80 nm BioReady Gold Nanospheres, cat. no. AUCR80-5M, nanoComposix, San Diego, CA, USA).

Antibodies and viruses: Anti-CCMV mouse monoclonal antibody (BAM-CCMV-29-81 used for LFA test line) was described previously [18,19,20]. Polyclonal rabbit antibodies (passively adsorbed to gold nanoparticles) and CCMV isolates were also published previously, including their production and purification protocols [18]. Polyclonal goat anti-rabbit IgG was used for the LFA control line (cat. no. 111-005-008, Jackson ImmunoResearch, Ely, UK); TMV, the lysine mutant, was kindly provided by Christina Wege, Institute for Biomaterials and Biomolecular Systems, Stuttgart, Germany [21]. CPMV (virus-like particle, cat. no. LES-P0001) was purchased from Leaf Expression Systems Ltd., Norwich, UK. Seeds of California Blackeye Number 5 (Vigna unguiculata) were purchased from Deaflora (cat. no. 97520, Werder, Germany).

2.2. Cultivation of the Host Plant Vigna unguiculata

Seeds of cowpea (Vigna unguiculata cv. “California Blackeye Number 5”) were soaked in tap water for 6 h at room temperature and subsequently wrapped in several layers of moist paper towels to initiate germination. The towels were placed in a Petri dish (5 cm in height) and sealed with parafilm to prevent drying. Germinated seeds were carefully transferred into the pre-wetted foam sponges of the growth system (iDOO, model ID-IG301, Eastvale, CA, USA), taking care not to damage the emerging radicle. The reservoir of the hydroponic unit was filled to its maximum volume (5 L) with water consisting of 50% tap water and 50% ultrapure water, supplemented with 0.1% (v/v) liquid hydroculture fertilizer. Seedlings were covered with a transparent plastic lid for an additional two days to maintain humidity. The water was replenished every two days, and plants were cultivated under a 16 h light and 8 h dark photoperiod at room temperature under the LED light of the hydroponic growth system.

2.3. Preparation of Virus-Free Plant Extract

Harvested leaves of 3-week-old (counting from the day of germination) virus-free plants were homogenized with running buffer (PBS with 1% Tween 20, pH 7.4) in a high-speed household blender. Approximately 40 g of leaves were homogenized in 250 mL of buffer (i.e., 1 g per 6 mL) to yield a plant extract. To prevent excessive foaming, a droplet of defoamer was added during the homogenization step. Afterwards, the crude extract was centrifuged at 15,000× g for 10 min and filtered using a 0.2 µm syringe filter. ProClin 300 was added to the plant extract at a final concentration of 0.05%, and aliquots were prepared and stored until further use at −20 °C.

2.4. Preparation of AuNP-Antibody Conjugate and Assembly of Lateral Flow Strip

Polyclonal rabbit antiserum against CCMV (BAM-CCMV-rab-pAb01) was purified as described in the previous work to obtain protein-G-purified polyclonal anti-CCMV antibodies (pAb) [18,19]. In brief, rabbit antiserum was micro-filtered (0.22 µm) and purified by Protein G affinity chromatography using a HiTrap Protein G HP (Cytiva, 29048581). Consequently, binding occurred with antibody binding buffer (8.7 mM Na_2_HPO_4_, 3.3 mM NaH_2_PO_4_, pH 7.2), and the polyclonal antibodies were eluted with antibody elution buffer (6 mM NaH_2_PO_4_, 6 mM H_3_PO_4_, pH 2.3) followed by neutralization with 100 mM NaOH to pH 7.4. For the preparation of the gold nanoparticle AuNP–pAb conjugate, 50 µL of 100 mM carbonate buffer (pH 10) was mixed with 10.5 µL (corresponding to 89.3 µg) of the polyclonal anti-CCMV antibody (pAb) solution (8.5 mg/mL in phosphate buffer, pH 7.4, containing 0.1% NaN_3_). Subsequently, 1000 µL of citrate-functionalized AuNPs (80 nm diameter, OD_520_ = 20) were added. The mixture was incubated for 3 h at room temperature on an overhead shaker to allow passive adsorption of the antibody (pAb) onto the nanoparticle surface. For blocking, 105 µL of a 10% (w/v) skim milk solution in 10 mM carbonate buffer (pH 10; sterile-filtered) was added, followed by an additional incubation for 45 min under gentle agitation. The conjugate suspension was stored at 4 °C until further use.

For the assembly of the lateral flow test strips, capture and control antibodies were deposited on nitrocellulose membranes using a sciFLEXARRAYER S3 piezoelectric spotter (Scienion AG, Berlin, Germany). The test line consisted of the monoclonal anti-CCMV antibody BAM-CCMV-29-81 (0.36 mg/mL in PBS) that was published previously [18,20], while the control line was spotted using a polyclonal goat anti-rabbit IgG. Spotting was performed with a drop volume of 320 pL and nine drops per spot at a pitch of 100 µm in both x- and y-directions with four parallel lines for each test and control zone. After spotting, membranes were immediately dried for 30 min in a vacuum oven at room temperature at 10 mbar. The membranes were then laminated onto adhesive backing cards without a conjugate or a sample pad. After application of the absorbent pad at the top, the lower portion of the backing card was trimmed off to complete the dipstick format. The assembled sheets were cut into individual strips of 4 mm width using a manual guillotine cutter and stored in sealed containers with desiccant until use.

2.5. Determination of Assay Sensitivity

Virus-free plant extracts as prepared in Section 2.3 were spiked with defined concentrations of CCMV (0, 2, 4, 8, 16, 32, 64, 124, 256, 512, 1024, and 2048 µg/L). Then, 100 µL of each sample was mixed in a non-binding microplate well with 3 µL of the AuNP–pAb conjugate as prepared in Section 2.4. Afterwards, test strips were immersed in the sample wells for 15 min before being imaged with the ChemoStar 2D Advanced Fluorescence and ECL imager (INTAS Science Imaging Instruments GmbH, Göttingen, Germany). Relative intensities of the control and test lines were determined using ImageJ (open-source software available here: https://imagej.net/ij/, Version 1.51, accessed on 23 April 2018). With this software, the differences in image intensities in the test line or control line regions from the background in the membrane strip region were calculated. Measurements were carried out in duplicate, the blank was measured in triplicate, and the relative standard deviation was determined. Relative intensities were then plotted against the corresponding CCMV concentrations. The LOD was determined using a logistic fit, where the average intensity of three blanks was multiplied by three times its standard deviation. LOD intensity values were translated to concentration values using a logistic calibration function.

2.6. Determination of Assay Specificity

Tobacco mosaic virus (TMV), cowpea mosaic virus-like particles (CPMV), and cowpea chlorotic mottle virus (CCMV) were diluted to a final concentration of 2000 µg/L in running buffer (PBS with 1% Tween 20, pH 7.4). Then, 100 µL of each sample, as well as 100 µL of running buffer as a negative control, was mixed in a microplate well with 3 µL of the AuNP–pAb conjugate as prepared in Section 2.4. Afterwards, test strips were immersed in the sample wells for 15 min before being imaged with the ChemoStar 2D Advanced Fluorescence and ECL imager (INTAS Science Imaging Instruments GmbH, Göttingen, Germany). Relative intensities of the control and test lines were determined using ImageJ (open-source software is available here: https://imagej.net/ij/, last accessed 15 December 2025). Measurements were carried out in duplicate, and the relative standard deviation was determined.

2.7. Leaf Sampling Protocol for Lateral Flow Analysis

For the field application of the simplified LFA presented in this work, the following sampling procedure (used in Section 3.4.1) is recommended:

Prior to sampling, 8 µL of AuNP-pAb conjugate solution (as prepared in Section 2.4) were pipetted onto the bottom side of the frit of a 2 mL polypropylene reactor and allowed to dry for 5 min at room temperature. Leaf disks of approximately 6 mg were excised from leaves using a handheld 6 mm hole punch and immediately transferred into the syringe. Per leaf, one disk was sampled. The disk was thoroughly ground with the plunger of the syringe until the frit color changed from red (coming from the AuNP-pAb conjugate) to green. Subsequently, 500 µL of running buffer (PBS containing 1% Tween 20, pH 7.4) was aspirated through the frit to reconstitute the conjugate and mix it with the crude extract. The extract was gently shaken for 15 s. Subsequently, approximately 100 µL (three droplets) were dispensed into a well of a non-binding 96-well microtiter plate. Finally, the simplified LFA strips without sample and conjugate pads (see Section 2.4) were then immersed in the prepared extracts for 15 min before visual and instrumental readout as described above in Section 2.5 and Section 2.6.

2.8. Monitoring of Vigna unguiculata After Infection with CCMV (Referenced in Section 3.4.2)

Mechanical inoculation of cowpea seedlings was performed 6 days after the start of cultivation as described in Section 2.2. First, frozen CCMV-infected cowpea leaves were mechanically crushed with a pestle in a 5 mL reaction vessel containing 3 mL of 50 mM sodium acetate buffer, pH 5.5. To 1 mL of this solution, a small amount of silicon carbide powder (600 mesh) was added. The suspension was vortexed immediately prior to application to ensure homogeneity. Then, the two primary leaves of each seedling (excluding the cotyledons) were inoculated by gently but firmly rubbing 50 µL suspension across the entire leaf surface (bottom and top). After 1 min of incubation time, the leaves were thoroughly rinsed with tap water. Starting on the third day after inoculation of the primary leaves, fresh samples were collected from a secondary leaf of the same plant on every weekday using a hole punch and immediately frozen at −80 °C. Ten days after inoculation, all eight samples per plant were crushed with a pestle in a 1.5 mL reaction vessel containing 500 µL running buffer. 100 µL were transferred to each well of a non-binding microtiter plate, and 2.5 µL AuNP-pAb conjugate was added. The test strips were imaged after 15 min and analyzed with the ChemoStar 2D Advanced Fluorescence and ECL imager (INTAS Science Imaging Instruments GmbH, Göttingen, Germany).

3. Results

3.1. Assay Format

For the development of lateral flow test strips targeting cowpea chlorotic mottle virus (CCMV), a monoclonal mouse antibody (BAM-CCMV-29-81) and a polyclonal rabbit antibody (pAb) were employed. Both antibodies have previously been described for CCMV quantification by ELISA [18,19]. In analogy to the optimized ELISA format, the lateral flow assay presented here utilizes the monoclonal BAM-CCMV-29-81 as a capture antibody at the test line. In the classical LFA configuration (Figure 1, right), the sample is first applied to a sample pad and migrates through a conjugate pad containing the AuNP-pAb conjugate before reaching the nitrocellulose membrane, where specific binding occurs at the test line. Detection of the analyte is achieved by accumulation of the AuNP-pAb conjugate, resulting in a visible red color signal at the respective line. Our simplified LFA format lacking both sample and conjugate pads is shown on the left side of Figure 1. Here, the AuNP-pAb conjugate is premixed with the sample prior to application onto the membrane.

An overview of the schematic assembly of the different lateral flow test strip designs is provided in Figure 2. The figure illustrates the arrangement of the individual components, including the nitrocellulose membrane and the presence or absence of sample and conjugate pads, highlighting the differences in design between the classic (left) and simplified (right) test strip formats. The premixed solution consisting of conjugate, buffer, and plant extract leads to a potentially more homogeneous mixture and a longer preincubation, which should be advantageous for assay performance.

A particular feature of this assay design is the absence of both sample and conjugate pads. This simplified “dipstick” format allows rapid assembly of the test strips without the need for labor-intensive impregnation and drying steps associated with conjugate pads. Particularly, the interfaces between the sample and conjugate pad, as well as the nitrocellulose membrane, are critical and can lead to unnecessary assay variation. Instead, the nitrocellulose membrane is directly immersed in a microtiter well or reaction tube containing a well-mixed blend of the sample, running buffer, and AuNP-pAb conjugate. To investigate a potential weakness of the “dipstick” design, namely that the AuNP-pAb conjugate remains on the membrane after flow-through and can thus increase the background and impair visual contrast, we quantified the membrane background in a line-free area (regions of interest, ROI) after developing the strips with three solutions: (i) running buffer, (ii) running buffer with AuNP-pAb conjugate, and (iii) raw plant matrix in running buffer. The background intensities were determined using a highly sensitive camera of the ChemoStar Imager.

As shown in Figure 3, the measured background intensities were comparable under all three conditions, with no significant increase in the presence of conjugate or plant matrix. Thus, remaining unbound nanoparticles do not measurably stain the nitrocellulose membrane, and the simplified format does not compromise either the visual contrast at the test line or the achievable limit of detection. Interestingly, all other components of the plant matrix appear to remain at the bottom of the nitrocellulose membrane, which can obviously take over the function of the sample pad. This underscores the premise that it is possible to dispense with a sample pad even when using complex matrices.

3.2. Determination of the Detection Limit

To evaluate the performance of the simplified LFA, crude extracts of virus-free host plants were spiked with defined concentrations of CCMV. The test strips were then applied, and both the test and control lines were examined. First, the visibility of the bands was assessed qualitatively by the naked eye. Subsequently, the signals were quantified using the high-sensitivity camera of the ChemoStar Imager (Version 0.5.74) by using standard focus light and a one-second shutter time. Finally, the resulting relative intensities of the test line regions were plotted against the spiked CCMV concentrations (Figure 4).

To determine the limit of detection (LoD), test strips corresponding to each concentration were imaged (shown in Figure S1), and the relative intensities of the test lines were quantified. A logistic fit was applied to the calibration data in the concentration range of 0–512 µg/L. Based on this analysis, the LoD in the plant matrix was calculated to be 3.5 µg/L, which corresponds to approximately 1 pmol/L or 100 attomole CCMV per 100 µL sample. This value is consistent with visual inspection of the test strips, as a concentration of 4 µg/L CCMV (the third test point) is still clearly discernible by the naked eye. The LoD of a previous ELISA performed with the same antibodies was around 0.25 µg/L. The assay exhibited a working range of approximately 10–100 µg/L, as inferred from the values presented in Figure 4. This also clearly shows that the nitrocellulose membrane can replace the sample pad and that it effectively retains the plant matrix of the raw extract.

3.3. Determination of Assay Selectivity

The cross-reactivity of the assay was assessed by spiking crude extracts of the virus-free host plant V. unguiculata with defined concentrations of cowpea mosaic virus-like particles (CPMV) and tobacco mosaic virus (TMV). CPMV was included due to the known susceptibility of V. unguiculata and its similar geographic distribution compared to CCMV [22], while a mutant of TMV was selected as a model plant virus. Subsequently, the LFA was applied in duplicate measurements, and relative intensities were quantified using a ChemoStar Imager. As shown in Figure 5, CPMV and TMV produced signal intensities comparable to the negative control (running buffer).

Thus, no cross-reactivity was observed even at high virus concentrations (2000 µg/L). Both the quantitative data and the visual inspection of the test strips confirm that the assay is highly specific for CCMV, at least for the tested samples.

3.4. LFA-Based Monitoring of CCMV Infection in V. unguiculata

3.4.1. Simulated Field Sampling with Endpoint CCMV Detection

A simulated field trial was conducted to distinguish infected from non-infected plants regardless of the occurrence of symptoms. For this purpose, plants at the two-leaf stage were mechanically inoculated with CCMV and tested with the LFA after 25 days according to different appearances. The sampling strategy for the field test is depicted in Figure 6.

Sample preparation is a critical, often underestimated step in sample analysis. To simplify it and combine it directly with the analysis, we developed a novel strategy for plant testing. The sampling method of monitored leaves is depicted in Figure 7.

Prior to sampling, the AuNP–pAb conjugate was preloaded onto the bottom side of the built-in filter of a 2 mL polypropylene syringe vessel as part of the device preparation and not necessarily immediately before field use. Accordingly, the syringe might be supplied preloaded with the dried conjugate as a ready-to-use component of a kit. For the experiments described here, 8 µL of the AuNP–pAb conjugate solution was applied to the filter and allowed to dry, as illustrated in Figure 7. Leaf material was collected as standardized disks of approximately 6 mg (see Supplementary Figure S2) using a handheld hole punch and transferred into the syringe vessel. The leaves were thoroughly ground using the syringe plunger, followed by aspiration of running buffer through the preloaded filter to resuspend the conjugate and extract the target analyte. The resulting crude extract was homogenized by gentle shaking, and approximately 100 µL were transferred to a well of a microtiter plate or another suitable vessel. For analysis, a simplified LFA strip (lacking both sample and conjugate pads) was directly immersed in the prepared extract. Notably, this format does not require a plastic cassette, which is commonly used in conventional LFAs [23].

Using this novel sampling method, infected and non-infected plants were examined, as shown in Figure 8.

Using the above sampling method, the time to achieve a result is 15 min when testing plants for systemic CCMV infection. The sensitivity of the assay enables the identification of symptomatic and asymptomatic infection courses. The specificity of the LFA is also shown and can, for example, rule out infection with CCMV in cases of chlorotic leaves or other similar symptoms. To prevent carryover between samples, a cleaning step with ethanol was performed after each use of the punch, followed by several punches through a paper towel (see Figure S3).

3.4.2. Pre-Symptomatic Detection of Systemic CCMV Infection

To evaluate whether the LFA detects systemic CCMV infection prior to the appearance of visible symptoms, plants at the two-leaf stage were mechanically inoculated with CCMV and subsequently monitored for ten consecutive days. Each day, one leaf at the three-leaf stage was sampled per plant and tested with the LFA. At the same time, plants were also photographed and monitored for the appearance of visible symptoms. A sample was classified as positive when the test line intensity exceeded the previously established LOD in the plant matrix. The plant cultivation and sampling strategy for plant monitoring is depicted in Figure 9.

Sampling secondary leaves after inoculating the primary leaves enables verification of systemic infection. Because secondary leaves develop slowly, sampling began three days post-inoculation, once sufficient leaf material was available.

Plant samples were taken daily from the same leaf of each plant and immediately frozen at −80 °C. Ten days after infection, all plant samples were examined in parallel using the LFA, as shown in Figure 10.

As part of the plant monitoring experiment, two plants (A and B; see Figure 10) were sampled and analyzed over a period of ten days following inoculation with CCMV. Plant A tested positive using the LFA on day 6 post-inoculation. On day 7, this plant began to develop visible symptoms, which became pronounced by day 10. In contrast, plant B did not exhibit any clear visual symptoms during the entire observation period, although it also tested positive on day 6 after infection.

A dilution series of samples collected on day 10 from plants A and B indicated some differences in viral load, which may account for the varying symptom severity (see Supplementary Figure S4). The LFA detection was possible for plants A and B at least up to a 1:1000 dilution. These results demonstrate that the LFA is suitable for monitoring systemic CCMV infections in plants and can detect infection prior to the appearance of visible symptoms, depending on the progression of infection and viral load.

4. Discussion

Our results show that a simplified lateral flow immunoassay without sample and conjugate pads can achieve sensitive plant virus detection while facilitating both manufacturing and use. Dipping the strip directly in a premixed well containing the sample and AuNP-pAb conjugate reduces variability associated with pad impregnation and drying. Furthermore, the conjugate mixed with the sample enables a relatively long and even incubation of the antibody with the antigen. In the conventional format, only the short flow time from the conjugate pad to the test line is available for antibody–antigen binding [24]. The analytical sensitivity (Figure 4) of our novel LFA for CCMV detection yielded an LOD of 3.5 µg/L, corresponding to 350 pg or 100 attomoles (~60 million virions) per 100 µL sample in only 15 min. An ELISA previously optimized with the same antibody pair and an assay duration of 5 h achieved a limit of detection of 0.25 µg/L [18].

For field applications and crop infection monitoring, we developed a straightforward method (Figure 7) that unifies sample preparation and test application. In this concept, the polypropylene frit of a syringe vessel serves three functions: conjugate reservoir, grinder, and coarse filter. This conveniently reduces handling steps, enables low-waste workflows, and supports sample-pooling strategies for surveillance, thanks to the small sample amount required (~6 mg plant tissue, corresponding to a single punched leaf disk). In our approach, the nitrocellulose membrane also serves several purposes: it replaces the sample pad used in conventional formats, draws the liquid upward through capillary forces, and serves as a surface for the heterogeneous immunoassay. We anticipate that this all-in-one approach will also benefit other dipstick immunoassays requiring on-site homogenization of complex matrices for analyte extraction (e.g., environmental samples).

The new method may have an obvious disadvantage compared to the usual LFA format, which is the lack of rinsing of the test membrane by pure running buffer due to the omission of the conjugate pad. This could lead to a higher background value due to the coloring of the plant matrix and the gold nanoparticle solution. Surprisingly, this effect was so weak that it was barely detectable (Figure 3).

The demonstrated performance and ease of use enabled rapid identification of infected plants in simulated field tests. While visible symptoms appeared about 7–10 days after inoculation, infection could be diagnosed as early as 6 days after CCMV inoculation using the proposed LFA format (Figure 9). This time advantage may permit management interventions, such as removing infected plants before vector-mediated transmission of an infectious dose to neighboring plants occurs. Furthermore, LFA-based analysis of plant samples, as shown in Figure 7, enables verification of visually ambiguous leaf tissue and helps discriminate true CCMV infections from chlorotic symptoms that would otherwise be misclassified as false positives.

In summary, the combination of a syringe-based sample preparation, a sample buffer vial, and conjugate delivery with a simplified LFA construction enables a compact, resource-efficient, and field-deployable assay without laboratory equipment such as readers, pipettes, grinders, filters, or centrifuges. The observed sensitivity and selectivity, combined with a practical sampling workflow, offer a viable route to routine on-site monitoring of CCMV and may provide new opportunities for the detection of other plant pathogens and targets when suitable antibody pairs of high quality are available [25].

5. Conclusions

By eliminating sample and conjugate pads and integrating sample preparation, conjugate reconstitution, and extraction into a syringe-based workflow, the assay reduces the complexity of the format while achieving a low limit of detection (3.5 µg/L in the plant matrix). The proposed approach is intended as a proof of concept rather than a finished commercial product. For example, further optimizations are needed to evaluate sustainability and validate long-term storage stability. Nevertheless, the combination of a mobile sampling method and a simplified test strip format offers a robust, field-deployable alternative to conventional LFAs.