Retention and infiltration of bacteria on a plant leaf driven by surface water evaporation

Evaporation is a fundamental process that can happen at various stages of preharvest and postharvest processing of leafy greens. Within an evaporating sessile droplet (i.e., a droplet that attached to a substrate) on a leaf surface, internal flows toward the contact line occur to compensate the high evaporation fluxes at the contact line region.^3,7–9 These flows can carry bacteria, if present, toward the leaf surface and at the location of contact lines¹⁰ and facilitate their access to the leaf interior.

Bacterial transport within an evaporating sessile droplet on a leaf surface can be due to several active and passive driving forces: (1) convective transport by the fluid flow,⁴ (2) diffusive transport due to Brownian motion¹¹ that is as a result of continuous bombardment from molecules of the surrounding fluid, (3) tactic motions or active swimming toward nutrient sources,²⁰ and (4) motile motions or the random runs and tumblings. Among these mechanisms, only the first one is purely induced by evaporation and can be a combination of capillary effects (driven by the surface tension of the liquid), thermocapillary effects (driven by the gradient of the surface tension of the liquid at the liquid-gas interface),^12,18 and flow due to the movement of the contact line. ¹⁷ 

A leaf surface is a complex environment that includes several microstructures such as trichomes, stomata, and grooves that can be attractive for bacteria.¹³ In addition to the availability of photosynthetic nutrients at the location of these microstructures (for example, stomata perform photosynthesis), they also serve as shelters for bacteria to avoid harsh environments and settle them at the leaf surface. As bacteria get hidden at the location of these microstructures, they cannot easily be washed away following typical sanitation practices, thus presenting a risk to the consumer. Since evaporation of water films and sessile droplets on plant leaf surfaces is a process that frequently happens as leafy greens move from field to fork, its role on the microbial contamination of the leafy greens needs to be better understood. The literature is scant on the specific effect of evaporation on bacterial retention at the leaf surface and their infiltration into leaf openings. This study intends to contribute to underlying mechanisms of this evaporation-driven retention and infiltration process. Due to the complexities of a leaf surface in terms of its hydrophobicity and roughness, artificial patterned surfaces fabricated out of polydimethylsiloxane (PDMS) are used here in conjunction with real leaves.

The objectives of this work were as follows: (1) To design and fabricate leaf surface surrogates out of PDMS patterned with three different common microstructures of stomata, trichomes, and grooves that are normally found on plant leaves. (2) To investigate how evaporation-driven flows, within a sessile droplet located on real leaves as well as fabricated patterned surfaces, lead to bacterial access to the surface microstructures. (3) To investigate the role of size and spacing of microstructures, and hydrophobicity of the surface, and bacterial concentration on bacterial deposition patterns. (4) To investigate how evaporation-driven movements of contact lines lead to bacterial infiltration into stomatal openings.

The experimental approach taken here includes fabrication and characterization of patterned PDMS surfaces, measurement of contact angles of sessile droplets on real leaves and the PDMS surfaces, and microscopic imaging of droplet (containing fluorescent bacteria/microparticles) evaporation on real leaves and the PDMS surfaces. Details of these experiments are discussed here.

The three common microstructures on the plant leaf surfaces were molded on the PDMS surfaces, as shown in Fig. 1. These microstructures include trichomes, stomata, and grooves whose dimensions were chosen based on the available microscopic imaging data.¹⁶ For each microstructure, two different sizes and spacings were fabricated. The PDMS surfaces were made in two different hydrophobicity levels within a range that many leafy vegetables sit (40° to 130°).¹⁴ 

Typical fabrication steps are shown in Fig. S1, available in the supplementary material. In step 1, photomasks were prepared for each type of microstructure. The patterns, shown in Fig. 1, were designed in L-Edit 15 (Mentor Graphics Corporation, OR, USA), and the CAD files (.gds format) were transferred to a Heidelberg mask writer (DWL2000, Heidelberg Instruments, Germany) to write the 12.7-cm chromium photomasks. After laser exposure, each photomask was developed and etched to remove the exposed layer of photoresist and uncovered chromium, respectively. Followed by a photoresist stripping, the masks were thoroughly rinsed and dried. In step 2, the molds required to pattern PDMS were made using a photolithography technique. ADEX sheets, dry-state negative photoresists, of various thicknesses were purchased from Integrated Micro Materials TM (TX, USA). The sheets were laminated on clean n-type one-sided silicon wafers of 100 mm diameter, using an ADEX hot-roll laminator (SKY-335R6, SKY-DSB, Ltd., Korea) at a roller velocity and a temperature of 0.3 m/min and 65 °C, respectively. Immediately after lamination, the wafers were baked at 65 °C for 30 min. Then, by using the photomasks fabricated in the previous step, the wafers were exposed to i-line UV light by a contact aligner (ABM, Inc., CA, USA) equipped with a short wavelength exclusion filter. The exposure dose for the ADEX sheets of 5 μm and 20 μm thickness was 90 mJ/cm² and 175 mJ/cm², respectively. This was followed by a postexposure bake at 85 °C for 10 min. The developing time for the ADEX sheets of 5 μm and 20 μm thickness was 5 min and 15 min, respectively, by using SU-8 developer. The developed devices were hard baked at 150 °C for 60 min. In order to avoid the PDMS sticking to the ADEX, a molecular layer of FOTS (a coating to make the surface hydrophobic) was deposited on the ADEX surfaces using a molecular vapor deposition machine (MVD-100, MVD TM).

In step 3, these ADEX devices were used as molds to pattern PDMS surfaces. Liquid PDMS (with a mass ratio of base-to-curing agent of 10:1) was vacuumed for 20 min to remove all trapped air bubbles. It was then poured onto the surface of the wafers and left to solidify at 65 °C for 120 min. The final patterned PDMS surfaces were placed on microscope cover-slips after applying oxygen plasma on the attaching surfaces of the glass and PDMS to improve their stickiness. The hydrophobocity of the final PDMS device was adjusted by depositing a layer of FOTS [(1H,1H,2H,2H-Perfluorooctyl)Trichlorosilane] (to make the surface hydrophobic) or APTMS [(3-aminopropyl)trimethoxysilane] (a coating to make the surface hydrophilic) using an MVD-100 machine.

After the fabrication of the PDMS devices, the size of the features was characterized by an optical microscope and an optical profilometer (NewView 7300, Zygo Corporation, CT, USA), as shown in Table I.

A contact angle goniometer (Rame-Hart 500, NJ, USA) was used to detect the variations of the contact angle of the droplet on various spinach leaves as well as the PDMS surfaces. For each experiment, a 1 ± 0.025 μl Milli-Q water droplet (without bacterial suspension) with a temperature of ∼23 °C was placed on the surface and the variation of the contact angle was imaged over evaporation time. Prior to deposition of the droplet on each surface, the surface was blown with a nitrogen gun for about 1 min to remove any possible dust and moisture. The contact angles were measured using ImageJ software with an uncertainty of ±0.5°.

To detect evaporation-driven bacterial collections on PDMS surfaces, a 1 ± 0.025 μl droplet of suspension of fluorescent E. coli RP437 cells (see Sec. S4 for the bacterial preparation procedure) was placed on each surface. Prior to the deposition of the droplet on each surface, the surface was blown with a nitrogen gun for about 1 min to remove any possible dust and moisture. The medium around the droplet was at a temperature of ∼23 °C and a relative humidity of ∼40%. The evolution of the contact line region was sequentially imaged using an inverted confocal microscope (Olympus IX71, Olympus Corporation, Japan).¹⁰ The initial droplet spreading radius and height on the APTMS-deposited flat surface were ∼921 μm and ∼645 μm, respectively. These values for the FOTS-deposited flat surface were ∼754 μm and ∼809 μm, respectively. When needed, the same procedure was applied using 1 μm fluorescent tracer particles (Bangs Laboratories, Inc., IN, USA) at a concentration of about 1 × 10⁸ particles/ml. Droplet evaporation, and bacterial deposition experiments on real plant leaves were done using an upright confocal microscope (Leica TCS SP5, IL, USA). Both confocal microscopy experiments were performed at 10× magnification and using a 488 nm argon laser.

This section presents first the variations of the contact angle of a sessile droplet on a real plant leaf and PDMS devices. Next, effects of the presence of stomata, grooves, and trichomes; their size and spacing; the hydrophobicity of the surface; and bacterial concentration on evaporation-driven retention and infiltration of plant leaves are discussed. Finally, the role of evaporation-driven movement of contact line in infiltration of bacteria into stomatal opening is elaborated.

The surface hydrophobicity of the plant leaves varies widely. For instance, lettuce leaves are hydrophilic with an initial contact angle of less than 45°, while spinach leaves are more hydrophobic, having a contact angle above 65°.¹⁴ Variation of the contact angle of sessile droplets during evaporation affects flow patterns within the droplet and thus transport of bacteria. Therefore, the variation of the contact angle of sessile droplets on leaf and patterned surfaces is analyzed here. Figure 2(a) presents the measured contact angle of 1 ± 0.025 μl sessile droplets at the surfaces of spinach leaves and flat PDMS surfaces covered with FOTS or APTMS. In general, the contact angle decreases during evaporation times almost linearly on both leaf and fabricated surfaces. Similar trends were found for the evaporation of sessile droplets on flat hot plates at various levels of hydrophobicity.¹⁹ This is because the contact line often pinned to the surface and removal of mass from the droplet led to reduction in its contact angle. The abaxial side of a spinach leaf is more hydrophobic than its adaxial surface.¹⁴ FOTS-deposited surfaces show closer surface characteristics to the abaxial side, while the APTMS-deposited surface better represents the adaxial side. See Fig. S3 for typical variations of the spreading radius, height, and volume of the evaporating sessile droplet on APTMS-deposited and FOTS-deposited flat PDMS surfaces, as well as respective vaporization rates.

Addition of the microstructures to the flat surfaces of the same hydrophobicity makes some changes in the variation of contact angle^1,21,22 during evaporation. For example, on APTMS-deposited PDMS patterned with trichomes, the contact angle decreased sharply during evaporation [Fig. 2(b)]. This is because trichomes enhance pinning of the droplet to the surface, and therefore, the contact angle decreases faster as the droplet evaporates. On stomata and grooves, the contact angle became somewhat constant after about 550 s due to the movement of the contact line over these microstructures. This constant contact angle trend was also seen on a flat surface [Fig. 2(a)]. On stomata, the contact line showed oscillations at these later times [Fig. 2(b)] due to a stick-slip behavior explained later.

The spacing of the patterned features highly affected the contact angle variation during evaporation [Fig. 2(c)]. Wider spacing reduced the contact angle. The narrow spacing probably brought the droplet closer to a Cassie state (i.e., droplet sits on top of the microstructures without any contact with the surface) and caused a superhydrophobic behavior. When spacing stayed the same, contact angles showed similar trends for two different sizes [Fig. 2(c)].

The oscillations seen in the contact angle on all three patterned surfaces after about 550 s of evaporation [Fig. 2(b)] are due to a stick-slip behavior of the contact line on the patterned surfaces.^2,6 Figure 2(d) shows a schematic of the underlying mechanisms of the stick-slip behavior of contact line on a surface patterned with trichomes. Initially, the contact line is pinned to the surface of the features. As the droplet evaporates, surface tension forces cause the contact angle to decrease. This reduction in the contact angle continues until it reaches a critical value⁵ at which the depinning forces generated from the surface tension of the evaporating droplet exceed the pinning forces [Fig. 2(d)], leading to the slipping of the contact line over the feature. This behavior can be observed in Fig. 2(e) that shows the evaporation of a 1 ± 0.025 μl sessile droplet on a FOTS-deposited PDMS surface patterned with trichomes of small size and wide spacing. The stick-slip behavior of the contact line can contribute to the bacterial infiltration into the leaf opening, as discussed in Sec. III C.

Figure 3(a) shows a ring formed on a spinach leaf surface after evaporation of a 0.5 ± 0.025 μl sessile droplet containing E. coli RP437 cells. When a sessile droplet containing bacteria evaporates on a leaf surface, it transports the bacterial cells close to the leaf surface and facilitates their access to the surface microstructures [Fig. 3(b)]. Figures 3(c) and 3(d) confirm the bacterial accumulation at the location of the contact line and into the stomata and grooves after evaporation of the sessile droplet. These findings imply the effects of evaporation-driven internal flows, shown in Fig. 3(b), in bacterial transport toward the contact line and close to the leaf surface. Similar patterns were also observed for 1 μm fluorescent beads (instead of bacteria) in a sessile droplet [Fig. 3(e)], highlighting the dominant role of passive transport by evaporation-driven internal flows (see also Sec. S2 for quantification of the flow velocity).

Plant leaves being complex, the underlying mechanisms of contamination can be studied more effectively by using fabricated surfaces with known hydrophobicity and roughness. Figure 4 shows the variations in the concentration of E. coli RP437 (fluorescence intensity represents bacterial concentration) during evaporation of sessile droplets, initially containing 10⁹ cells/ml on APTMS-deposited surfaces. On a flat surface [Fig. 4(a) (Multimedia view)], the highest concentration of bacteria is at the contact lines (the sharp line of the fluorescence intensity). In the presence of stomata [Fig. 4(b) (Multimedia view)], bacteria are also collected within the features. Stomatal pores located at the contact line region contain the highest concentration of bacteria after evaporation [see also Fig. 3(c)]. Trichomes play the role of micropillars in front of internal flows that can trap bacteria. Figure 4(c) (Multimedia view) shows how bacterial cells are collected around trichomes during evaporation. At the contact line, more bacteria are collected around the trichomes, which is due to the bacterial transport by evaporation-driven flows toward the contact line. Bacteria are also deposited within the grooves [Fig. 4(d) (Multimedia view)]. Grooves located at the contact lines contained a much higher concentration of bacteria after evaporation, as is highlighted by the fluorescent intensity profile. A comparison between the current findings on the PDMS surfaces patterned with stomata and grooves, with those obtained on the real leaves [Figs. 3(c) and 3(d)], highlights the applicability of the fabricated surfaces in mimicking plant surface microstructures. Once bacteria are located inside or around stomata, they can further infiltrate into the leaf interior during longer times by chemotaxis toward the concentration gradients of photosynthetic sugars.¹³ 

Stomata are the main natural routes for infiltration of bacteria into plant leaves. The effect of their size and spacing on the bacteria infiltration is shown in Figs. 5(a)–5(c) (Multimedia view). Stomatal density on the adaxial side of a leaf is lower than its abaxial side. Therefore, patterned surfaces with stomatal features of wide spacing [Fig. 5(a) (Multimedia view)] and narrow [Fig. 5(b) (Multimedia view)] spacing can represent adaxial and abaxial sides of a leaf, respectively. An increase in the stomatal density led bacteria to mainly accumulate at the location of the contact line. The presence of stomata can create upward flows away from stomatal pores that can transport unattached bacteria out of the pore. When the stomatal density increases, these flows get stronger. A combination of these upward flows and the evaporation-driven downward and radial flows close to the leaf surface leads bacteria to mainly transport through rows between stomatal pores toward the contact line. Therefore, they are rarely trapped inside the high-density stomatal pores [Fig. 5(b) (Multimedia view)] and mainly accumulate at the contact line region.

Small stomatal features with wide spacing [Fig. 5(c) (Multimedia view)] can represent partially closed stomata on the adaxial side of a leaf at which the stomatal density is lower.¹⁵ Comparing Figs. 5(a) and 5(c) (Multimedia view), a wider stomatal pore seems to trap more bacteria per pore. The bacterial infiltration into stomatal openings during evaporation is concentration-dependent. Figure 5(d) (Multimedia view) shows the effect of 1 order of magnitude decrease in the bacterial concentration [compared with what is used in Fig. 5(a) (Multimedia view)] on their infiltration into stomatal pores. Obviously, the infiltration (presence of bacteria inside the pores as represented by fluorescence intensity) is more noticeable when the bacterial concentration in the droplet is higher. Surface hydrophobicity can also affect the bacterial accumulation patterns within an evaporating droplet. A more hydrophobic surface [Fig. 5(e) (Multimedia view), FOTS-deposited] drastically reduced infiltration of bacteria within the stomatal pore area compared to a hydrophilic surface [Fig. 5(a) (Multimedia view), APTMS-deposited]. The hydrophilic nature of the surface can keep the bacteria closer to the leaf surface. Therefore, the chance of bacteria to infiltrate the stomatal pores on a hydrophilic surface [Fig. 5(a) (Multimedia view)] is higher than that on a more hydrophobic surface [Fig. 5(e) (Multimedia view)].

Video microscopy [Fig. 6(a) (Multimedia view)] reveals that mainly on a more hydrophobic surface (FOTS-deposited), when the evaporation rate is high enough, the contact line sticks and slips over the microstructures. Here, instead of bacteria, fluorescent microparticles were used to show that this mechanism can lead to particle attachment to the microstructure even without any active attachment means, such as flagella that wild-type bacteria have. Evaporation-driven flows transport microparticles toward the contact line.⁴ The receding movement of the contact line over stomatal pores, as a result of evaporation, leads to microparticle deposition at the edges or within the pores. This observation can be explained using a schematic shown in Fig. 6(b): in the first (stick) stage, the contact line sticks to the edge of the stomatal pore. Evaporation-driven internal flows transport bacteria/microparticles into the stuck region where they can attach to the surface and edges. Meanwhile, the surface tension forces tend to pull the liquid surface in an opposite direction (toward the droplet centerline), leading to a local reduction in the contact angle at the stuck edge. This smaller contact angle creates a higher evaporation flux at the stuck edge (as it acts like a hydrophilic surface) and a stronger microparticle transport into the stuck region. As the droplet evaporates, the surface tension forces cause the apparent contact angle of the droplet to decrease to a critical value at which the depinning forces dominate the pinning forces [Fig. 2(d)]. At this second (slip) stage [Fig. 6(b)], the contact line slips over the stomatal pore and leaves the attached microparticles/bacteria at the stomatal edge.

Evaporation of sessile droplets containing bacterial suspensions on plant leaves was studied using confocal microscopy. Due to complexity of plant leaves in terms of variability in the surface roughness and hydrophobicity, fabricated PDMS surfaces were used to further understand the underlying mechanisms of microbial retention and infiltration during evaporation. Droplet evaporation experiments on plant leaves showed that evaporation-driven flows can transport bacteria close to the leaf surface and facilitate their access to the microstructures, such as trichomes, stomata, and grooves, leading to their significant infiltration into the stomatal opening. Larger size and wider spacing of the micropores, and a more hydrophilic surface, led bacteria to spread more on the droplet base area and infiltrate into more stomata. The infiltration was more noticeable when the bacterial concentration in the droplet was higher. During evaporation of a droplet on stomatal pores, the contact line may stick to the pore edges and slip over them. Sticking of the contact line to the stomatal pores increases the time scale at which evaporation-driven internal flows can transport bacteria into the stomata that facilities their infiltration.

See the supplementary material for the fabrication steps, the estimated flow within an evaporating sessile droplet, and the bacterial preparation procedure.

This work was supported under Grant No. 2014-70003-22357 by the USDA National Institute of Food and Agriculture. Microfabrications were partially supported by a Spencer award. The microfabrications were done at Cornell Nanoscale Science and Technology Facility (CNF). Special thanks to Dr. Beth Rhoades at CNF for all her guidance during microfabrications and experimentation steps. Also, the authors gratefully thank Professor Mingming Wu in the Biological and Environmental Engineering (BEE) department of Cornell University for guidance on the microfabrications and experimentations, as well as providing bacteria. Bacterial preparation for each experiment was done in Professor Ludmilla Aristilde’s lab in the BEE department of Cornell University. The authors acknowledge Mina Solhtalab, Ph.D. candidate at BEE department of Cornell University, for all her contributions in bacterial preparation used in this work. The confocal microscopy imaging of plant leaves was done in the Plant Cell Imaging Center (PCIC) at the Boyce Thompson Institute (BTI) of Cornell University with the help of Dr. Mamta Srivastava.