Artificial surfaces are routinely used instead of leaves to enable a reductionist approach in phyllosphere microbiology, the study of microorganisms residing on plant leaf surfaces . For instance, flat surfaces such as nutrient agar , enable the influence of nutrient supply on micro organisms to be investigated . In contrast microstructured surfaces, su ch as isolated leaf cuticles or reconstituted leaf waxes enable the influence of physic ochemical properties to be investigated . However, interest in replica leaf surfaces as an artificial surface is growing . As replica surfaces offer an improved representa tion of the complex topography of leaf surfaces. The use of r eplica leaf surfaces has to date primarily been focused on replicating the superhydrophobic surfaces of leaves. Whereas i n this paper , we investigate potential replica surface materials for phyll osphere microbiology studies . Using a test pattern, we in vestigated the resolution, the degradation characteristics in environmental conditions, surface energy , and bacterial survival characteristics for each potential replica material . Our r esults indicat e that PDMS is the most suitable material for producing replica leaf surfaces . Due to the high resolution achieved through replica molding, extended stability, hydrophobic properties , and bacterial survival characteristics comparable to isolated leaf cutic les. Our experiments highlight the importance of considering the inherent material properties , when selecting a replica leaf surface for phyllosphere microbiology studies . A replica leaf produced in PDMS offer s a control surface that can be used for invest igating microbe -microbe and microbe -plant interactions in the phyllosphere . Thus, in turn enabling mitigation strategies against pathogens to either the plant host or humans to be developed. 3 1 Introduction Many microorganisms thrive on plants . They resid e either permanently or temporarily in the plant environment. Furthermore, microorganisms contribute to the health of the plant host. These microorganisms live in three interconnected compartments: the spermosphere, rhizosphere, and phyllosphere.1-6 To date research has primarily be en focused on the rhizosphere. T he rhizosphere is the microbial habitat in the vicinity of the pla nt roots, which is influenced by the products secreted by the roots .6, 7 However, recently phyllosphere microbiology , the study of microorganisms which reside on plant leaf surfaces, has gained increasing interest. This increase is attributed to growing interest in the role that microorganisms in the phyllosphere , have on the health of the plant host. Microorganisms in the phyllosphere are in direct contact with the plant cuticle. Where the plant cuticle , a protective waxy film that coats the leaves of plants and prevents pathogenic attacks against the plant host .8 In addition, the cuticle prevents water, ion , and nutrient l oss.9 Leafy greens are exposed to a range of potential contamination sources during g rowth on a farm , resulting in unwanted microorganisms in the phyllosphere . This is of importance for produce grown for human consumption, such as lettuce, rhubarb, and parsley .4-7, 10 Potential contamination sources include : irrigation water, soil, fertiliser, farm workers , and equipment used around a farm .11, 12 Unwanted contamination can result in diseases detrimental to plant health or produce unwanted pathogens that are harmful to humans . Furthermore, lea fy greens are often consumed raw or with minimal processing . Which does not remove or kill unwanted contamination .4, 10 -14 In some cases, contamination can lead to outbreaks that can cause severe illnesses .15-17 Unwanted leaf contaminat ion is of an increased concern with increasing produce demand, large -scale production , and distribution. Consequently, further studies to understand phyllosphere microbiology are imperative for develop ing mitigation strategies . For instance, microbe -microbe and microbe -plant intera ctions need to be studied in greater detail to determine said mitigation strategies. Potential mitigation strategies may include : introducing other microorganisms to pr event against pathogens detrimental to either the plant host or humans ; or developing different clea ning protocols of leafy greens.17, 18 The use of artificial surfaces is common practice in phyllosphere microbiology . Such surfaces are often utilised to provide an insight into microorganism communities in the phyllosphere of leafy greens . Artificial surfaces are used instead of a living leaf to enable a reductionist approach. Such an approach , allows one to identi fy the impa ct of individual factors on microorganism functio ning and viability .13 For instance, an artificial surface provides a controlled environment to study contamination of leaf y greens .19, 20 Artificial s urfaces can be classified as either flat or microstructure d, due to th eir lateral heterog eneity . 4 (1) Flat surfaces. A flat surface is defined as a surface that is laterally homogenous. In general, a flat surface will have no lateral chemical or biological heterogeneity at the time of inoculation with microorganisms . Commonly used flat surface s include nutrient agar or inert surfaces (for example, metal, plastic , and glass ). Nutrient agar is commonly used to investigate the influence of different nutrient compositions on microorganism interactions and colonisations . For example, Jacobs et al. investigated the role of pigmentation, ultraviolet radiation tolerance , and leaf colonisation strategies in epiphytic survival using nutrient agar .21 Inert flat surfaces on the other hand , are commonly utilised for the investigation of attachment processes of microorganisms . For example, Rivas et al. observed variati on in surface attachment amongst strains of Shiga toxigenic Escherichia coli (E. coli ) on stainless steel surfaces .22 (2) Microstructured surfaces. Microstructured surface s are defined as surface s which are laterally heterogeneous . Furthermore, m icrostructured surfaces generally have no lateral chemical or biological heterogen eity at the time of inoculation. A microstructured surface is more representative of the surface of the leaf , as the cuticle of a leaf is physically (and chemically) laterally heterogeneous.23 Commonly used m icrostructured surfaces include recons tituted leaf wax, leaf peels, isolated leaf cuticles , and microfabricated surface s.24-26 For instance, Remus -Emsermann et al. examined the permeability of fructose through isolated poplar ( Populus × canescens ) cuticles , to assist in explaining microorganism growth patterns in the phyllosphere .27 Although microstructured surfaces are suitable for their respective application s, they do not entirely represent the complex nature of the topography of plant leaves .28, 29 Recent studies have utilise d double -casting protocols to overcome this limitation. As such replica leaf surfaces have been produced in agarose, dental wax , or polydimethylsiloxane (PDMS) .28-31 To date leaf surfaces have been predominately reproduced for their self -cleaning properties .32-34 In phyllosphere microbiology, studies are beginning to appear using microfabricated replica leaf surfaces . In one example, Zhang et al. produced replica spinach leaf surfaces in agarose . They investigated the interaction of E. coli on flat agarose and agarose replica leaf surfaces .31 However, studies to date, including the aforementioned did not compare the potential influence of the replica leaf material have on microorganism viability and behavior. In general, m aterials used as replica leaf sur faces to date have not been fully investigated for their use in phyllosphere microbiology studies . As the inherent properties of a material can influence microorganism behaviour , it is imperative that the suitability of a material as a replica leaf surface is characterized comprehensively . Investigations should include: (1) the resolution and degradation characteristics of the replica leaf surfaces . To determine if the replica can suitably mimic the co mplex topography of a leaf surface and maintain the topography throughout an experiment. (2) The 5 hydrophobicity of a replica leaf material and how representative this is of a leaf surface . As hydrophobicity influences the attachment processes of microorgan isms. We selected three commonly -used biocompatible materials to investigate their suitability for replica leaf surface materials , for phyllosphere microbiology studies . (1) A garose. Agarose has been used as a replica leaf surface, has well established mic rofabrication protocols , and is routinely used in phyllosphere microbiology studies .13, 31, 35 (2) PDMS. PDMS is routinely used in biological applications, such as the foundation of lab -on-a-chip devices and in bioi mprinting .36-42 (3) Gelatin. Gelati n, was initially used as a gelling agent for microbiology growth media, until it was succeeded by agar.43 In addition, well establi shed microfabrication protocols exist for gelatin .44 In total, t hree concentrations of each material w ere used in our comparison . For our investigation , we used a test pattern comprised of regular circular pillars . The pillars were designed to have a height of 5 µm and a diameter of 15 µm to mimic microfeatures found on leaves – trichomes, stomata, and grooves .20 For eac h material we measured : (1) O ptical surface profiles . (2) The degradation of the materials over three days . (3) The hydrophobic properties aga inst two generic isolated leaf cuticles ( Citrus × aurantium , and Populus × canescens ). In addition, t o investigate the biological suitability of each material we used a model leaf colonising bacterium Pantoea agglomerans 299R . We compared the bacteria surv ival results of the materials, to bacterium on isolated Citrus × aurantium cuticles. 6 2 Materials and Methods An overview of the fabrication protocols to produce the patterned materials is provided in Fig. 1, with accompanying procedures explained in the fol lowing sub-sections. In brief, a negative -tone photoresist mold master was fabricated using standard soft -lithography processes . The mold master comprised of an array of circular pillars, with a height of 5 µm and a diameter of 15 µm. From the mold master the polydimethylsiloxane (PDMS ) templates were produced . In turn, the PDMS template was used to fabricate patterned agarose, PDMS , and gelatin substrates. The patterned materials were then used in our investigation towards finding a suitable leaf surface r eplica material. In this process the mold master and PDMS template are representative of a living leaf surface and leaf imprint, respectively . Whereas, t he patterned materials are representative of a replica leaf 28-31. We selected PDMS as our template material as PDMS is well-established in microfabrication . In addition, PDMS can exhibit nano scale pattern resolution .41, 42 2.1 Mold Master Fabrication For the mo ld master substrat e a 4” prime grade silicon wafer was used. The wafer was dehydrated at 185 C overnight in an oven to improve resist adhesion . The wafer was then removed from the oven and left to cool to room temperature. The wafer was subsequently cleaned in oxygen plasm a for 10 minutes. ADEX05 (DJ MicroLaminates), an epoxy dry film photoresist, was then processed to produce a mold master.39 Prior to casting the PDMS template, the mold master was treated with Trichloro(1H,1H,2H,2H -perfluorooctyl)silane (448931 , Sigma -Aldrich) for two hours . This treatment was undertaken to facilitate the remo val of the PDMS template. 2.2 PDMS Template Fabrication The PDMS template was produce d using standard replica moulding techniques (Fig. 1a).45 The PDMS (Sylgard 184, Dow Corning) was prepared at a ratio of 10:1 w/w (base to curing agent). The base and curing agent were thoroughly mixed together , and then degassed in a vacuum desiccator (Z119016 , Sigma -Aldrich) until no bubbles r emained. Following this, the PDMS was poured onto the photoresist mold master and degassed again . Once no bubbles remained, the PDMS and mold master were placed on a hot plate for two hours at 80 ℃ to allow the PDMS to set. Once set, the PDMS template was carefully removed from the mold master. The PDMS template was then placed on a hotplate for a further two hours at 80 ℃ - to improve the durability of the template. Several templates were fabricated to produce enough patterned samples for all experiments. All patterned substrates were fabricated to a n overall height of 3 mm . This was achieved by usi ng rings with an internal diameter of 66 mm and filling the ring to the 3 mm line. From the patterned substrates 7 smaller samples were taken using a cork borer (Usbeck, Germany) , with a n internal diameter of either 7.8 or 11.5 mm . 2.3 Patterned Agarose Fabrica tion In preparation of casting the patterned agarose, the PDMS template s were placed under vacuum for two hours. The agarose (Agarose Low EEO, A0576, Sigma -Aldrich) was added to phosphate buffered saline (PBS, P4417, Sigma -Aldrich) to produce a concen tration of 2.5, 5 , or 7.5 % w/v . The agarose solution was then placed on a hot plate at 200 ℃ , until the agarose was fully dissolved – approximately 15 minutes. Once the powdered agarose was fully dissolved, the solution was then poured onto the degassed PDMS template . This stack was then placed in a vacuum desiccator (Z119016 , Sigma - Aldrich) for ten minute s with an open outlet valve (Fig. 1b(i)). Following this, the stack was left at room temperature to allow for the agarose to set for an hour. Once set , the patterned agarose was carefully peeled off the PDMS template .35, 46 2.4 Patterned PDMS Fabrication In preparation for casting the patterned PDMS, the PDMS template was treated with 0.1% w/v hydrox ypropylmethylcellulose (H8384, Sigma -Aldrich) in a phosphate buffer saline (PBS, P4417, Sigma -Aldrich) for 10 minutes . The patterned PDMS was prepared at a ratio of 5:1, 10:1, or 20:1 w/w (base to curing agent). The b ase and curing agent were thoroughly mi xed together , and the mixture was then degassed until no bubble s remained (Fig. 1b(ii)). The PDMS was then poured onto the PDMS template and degassed again . Once no bubbles remained, th is stack was then placed on a hot plate for two hours at 80 ℃ to allow the patterned PDMS to set. Once set, the patterned PDMS was carefully removed from the PDMS template .47, 48 2.5 Patterned Gelatin Fabrication Gelatin from por cine skin ( gel strength 300, G2500, Sigma -Aldrich), was added to PBS (P4417, Sigma -Aldrich) to produce a concentration of 10, 12.5, or 17.5 % w/v. The gelatin solution was then placed on a hot plate at 50 °C until the gelatin was fully dissolved – approximately 30 minutes (Fig. 1b(iii)). The microbial transglutamina se (mTG, Ajinomoto Co., Inc., activity of approximately 1000 U g-1) solution was prepared in 1 ml of PBS . The mTG solution was prepared to a final concentration of 10 U of mTG per one gram of gelatin . The mTG solution was thoroughly mixed to ensure the mTG was fully dissolved . Once dissolved, the mTG solution was added to the gelatin solution and quickly mixed together. Once thoroughly mixed, the gelatin -mTG solution was immediately poured over the PDMS template. After pouring, the gelatin on the PDMS templ ate was left at room temperature for five minutes to stabilise . The stack was then placed into an oven at 37 °C for four hours to set the 8 gelatin . To enable the gelatin to be readily peeled off the PDMS template, the stack was removed from the oven and pl aced in a fridge at 4 °C for 30 minutes. The patterned gelatin was then carefully peeled off the PDMS template .44 2.6 Optical Profiles All optical profiles for the patterned substrates were obtained within 30 minutes after peeling from the PDMS template . This was done to minimise potential effects from degradation due to the ambient conditions in the laboratory. All 3D optical profiles were obtained using a Profilm3D optical profilometer (Filmetrics Inc., USA) , equipped with a 20 × objective (CF Plan 20×/0.40 DI, Nikon). For agarose and gelatin , the high sidewalls of the pillars made imaging difficult due to light scat tering. To correct for this, the data was processed using the inbuilt remove outliers function in the Profilm 3D software 2018 (ver. 3.2.7.2, Filmetrics Inc., USA) . For the remove outliers function, the invalid pixels filled in method was u sed, and with maximum slope set to 10. During post processing, all images were also filtered using a 3-point level function in Profilm3D software to compensate for non -level placement of the substrates . 2.7 Atomic Force Microscopy Scan s Atomic force microscopy (AFM) scans were obtained for the mold master , PDMS template , and patterned PDMS. Due to the patterned agarose and gelatin degrading during the required imagining period, the AFM tip would lose contact. As a result, AFM images were not possible for agarose and gelatin. All images were taken using a Digital Instruments Dimension 3100 (Vecco, USA) equipped with TAP 300-G tips (BudgetSensors, USA) operating in tapping mode. All AFM scans were analysed using Gwyddion (Version 2.49). 2.8 Degradation Measurement Method Two condition s were examined to test the degradation of the patterned materials: (1) 30 C at a relative humidity of 25%, and (2) 30 C at a relative humidity of 75%. A temperature of 30 C was selected as it sustains bacterial life. Whereas , the humidity levels were s elected to : (1) mimic dry conditions, similar to a climate -controlled laboratory ; and (2) a higher humidity that would slow the degradation of the patterned samples (prepared as detailed in the following sub -section) . Five samples per the three different c oncentrations for each material were used. Each sample had an initial diameter of 11.8 mm. Weight measurements were taken at: 0, 2, 4, 6, 12, 24, 48, and 72 h. A weightless percentage measure was then determined for each of the time points. Results are pre sented as mean ± SEM (standard error mean). 9 2.9 Humidity The degradation of agarose and gelatin is undesirable for investigating microorganisms in the phyllosphere. Consequently, an environment was sought to minimise the degradation of the materials . In additi on, the environment must be favourable for microorganisms, and minimise the potential swelling of the materials. To achieve this a saturated salt solution was prepared by dissolving 72 g of sodium chloride (71382, Sigma -Aldrich) in 200 ml of deionised wate r. This solution was then kept in an open top container alongside samples for degradation and bacteria recovery experiments, inside an airtight container. The airtight container was placed in an oven at 30 °C, which resulted in a constant relative humidity of 75 % (see Supplementary Fig. S1 for humidity data) .49-51 2.10 Contact Angle Measurement with Water Method All contact angle measurements were undertaken with a CAM200 (KSV Instruments Ltd, Finland) , integrated with K SV CAM Optical Contact Angle and Pendant Drop Surface Tension Software ( ver. 4.01, KSV Instruments Ltd, Finland). Flat samples were compared against patterned samples for each material at the three selected concentrations . Five samples were measured for ea ch concentration. All samples had a diameter of 11.8 mm . This diameter was selected to minimise the potential occurrence of edge effects on the water droplets . For agarose , which is considerably hydrophilic, all results presented are for droplets with a vo lume of less than 40 µl. Whereas f or PDMS and gelatin, water droplets with a volume less than 60 µl were analysed . Prior to the contact angle measurements all samples were dried with dry nitrogen gas. This was done to minimise any potential effects from surface moist ure. In addition , experiment s were conducted within an hour of peeling the material away from the PDMS template . Deionised water was used to determine the surface energy of the material s. Results are presented as mean ± SEM (standard error mean) . For statistical analysis, Student ’s t- test or ANOVA was performed using GraphPad Prism 7 (GraphPad Software, USA). P values less than 0.05 were considered significant (*P<0.05, **P<0.01, ***P<0.001 , and ****P<0.0001). 2.11 Bacteria Culture Protocol Pantoea ag glomerans 299R , a model leaf colonising bacterium isolated from a healthy leaf of a pear tree, was grown overnight on nutrient agar plates ( 13 gL-1 Lysogency broth a nd 15 gL-1 bacteriological Agar, Oxoid) at 30 ˚C.52 The P. agglomerans was then harvested using a sterile inocul ation loo and resuspended in 5 ml of sterile phosphate buffer (8 gL-1 NaCl (LabServ), 0.2 gL-1 KCl (LabServ), 1.44 gL-1 Na2HPO 4-GPR (AnalaR ), 0.24 gL-1 KH 2PO 4 (AnalaR), pH 7.4). Following this, b acteria were washed by centrifugation at 1150 RCF for five m inutes at 10 C. The supernatant was discarded, and the bacteria was suspended in fresh phosphate buffer to a n OD 600 nm of 0.2 , corresponding to approximately 2 x 108 bacteria per mL. 10 2.12 Bacteria Viability Protocol For bacteria viability experiments samples w ith a diameter of 7.8 mm were used . Five samples were taken per time point. All samples were sterilised for 15 minutes with u ltra-violet sterilisation. Then 100 µL of bacteria l solution ( ~2 x 107 bacteria ) was inoculated to the patterned sample s, using a n air brush (KKmoon T-180 Airbrush , China ) at 1×105 Pa.53 Patterned samples coated with bacteria were then incubated at 30 ˚C at a relative humidity of 75%. Five samples were randomly selected per measurement time point – 0, 2, 4, 6, 12, 24, 48, and 72 h. Each individ ual sample was suspended in 1 mL of fresh phosphate buffer in a 1.5 mL Eppendorf tube. Individual s amples were then placed in a Bead Ruptor 24 (Omni Internatio nal Inc., USA ) at 2.6 ms-1 for two sets of five minutes , with a five minute interval. Following this, t he samples were placed in an ultrasonic bath (E Easy Elmasonic, Elma Schmidbauer GmbH , Germany) for five minutes . For each sample 100 µL of bacterial solution was recovered .54, 55 The recovered solution was then p lated onto nutrient agar by undertaking serial dilution s using PBS. At each dilution step 10 µL aliquots were taken. W ith a final dilution of 10-6 being plated. Colony -forming units (CFU) were expressed as bacterial cell density per unit area for each sample (7.8 mm diameter). The cell density was normalised to the initial number of cells after inoculation . This was undertaken t o enable a comparison between the survival curves of the bacterial colonies on the different materials. Results are presented as mean ± SEM (standard error mean). 11 Fig. 1. Schematic overview of the fabrication protocols. (a) Protocol used to produce the PDMS templates. (b) Protocols used to produce the patterned: ( i) agarose, ( ii) PDMS , and ( iii) gelatin. 12 3 Results and Discussions 3.1 Pattern Resolution The microstructures found on lea ves are dependent on the plant species and can range in complexity . Microstructures found on leaves include groves to stomata (pores) to the more complex stru cture of trichomes (leaf hairs). The ability to sufficiently replicate the surface topography of a living leaf into a replica , is important for produc ing r eplica leaf surface that can be used as a control surface in phyllosphere microbiology. In addition, inherent damage due to the replication process could produce inaccuracies of the replica leaf surface . This is important as the topography of the leaf surface has the ability to influence colonisation and attachment behaviours of microorganisms .4, 20 To investigate this with our selected materials we used a regular test pattern . The test pattern consisted of circular pillars with a height of 5 µm and a diameter of 15 µm. Such dimensions are comparable to the features found on leaf surfaces – trichome, stomata , and grooves. For example, Pelargonium zontal stomata have a length and width of 16 µm by 9 µm .56 In contrast, t richomes on a tomato leaf have a width of approximately 17 µm at the ir base.20 Comparisons of the patterned materials were undertaken using 3D optical profiles (Fig. 2). Optical profiles were selected over more common ly used techniques to compare microstructure resolution , such as atomic force microscope (AFM) and sca nning electron microscopy (SEM). This was due to the inherent properties of agarose and gelatin influencing the ability to obtain high -resolution images . The ability to use AFM imaging was hindered by the softness and quick degradation of both agarose and gelatin. As such, when investigating the suitability of AFM imaging, we regularly observed the AFM tip losing contact during imagining . On the other hand, for SEM imaging a conductive coating would be required . As both agarose and gelatin are non -conductive. To produce a conductive coating the sa mple s would have to be placed under vacuum. A short three minute vacuum was tested which resulted in a noticeable structural degradation of both materials ( data not shown ). Obtaining an optical profile instead require s no modification to be undertaken on the materials , and scans can be completed in approximately three minutes - mitigating any affects from degradation (see section 2.6 Optical Profiles for more information ). To prove the validity of optical profilometr y for our materials we compared our results to AFM scans undertaken on the mold master, PDMS template, and the patterned PDMS (see Supplementary Information S2 and Table S 1). The measurements obtained using the AFM and optical profilometer are in agreeance (Table 1 and Table S 1). For example, the height of the mold master was measured to be 4.50 ± 0.03 µm using the optical profilometer - which lies in the rang e measured by the AFM of 4.44 ± 0.16 µm. Furthermore, the measured widths were 14.64 ± 0.47 µm and 14.94 ± 0.21 µm for 13 images taken by the AFM and optical profilometer, respectively. In consequence, this indicat es no significant difference between 3D profi les taken by either AFM or an optical profilometer . A summary of optical profilometry measurements of the height and width of the pillars is presented in Table 1 – or in the case of the PDMS template , the depth and width of the well. Twenty pillars were measured for each dimension measurement. In the case of agarose , the pillars were smaller by 1.54 ± 0.57 µm (in height) and 4.93 ± 0.25 µm (in width) in comparison to the mold master (Table 1) . In addition , a larger variation in measured heights for agarose was observed . We conjectured this variation was most likely a result of non- uniform shrinkage occurring while the agarose sets during fabrication .57 In addition, pillars produced in gelatin were also smaller. With the height and width of the pillars in gelatin 0.61 ± 0.03 µm and 1.14 ± 0.27 µm smaller than the mold master, respectively (Table 1) . We conjecture d this was due to the gelatin degrading while setting, during fabrication . In addition, we observed that pillars regularly broke when peeling the gelatin off the PDMS template. We minimised the occurrence of the pillars breaking by following the recommendations made by Paguirigan and Beebe . In which the gelatin and PDMS templat e were placed in the fridge prior to peeling off the gelatin .44 Trichomes (leaf hairs) would more than likely not withstand being peeled off the PDMS template. This problem is compounded by t richomes being quite intricate and considerably fragile .20 In summary, patterned PDMS was more representative of the patterned pillar microstructures used as mold master . Dimensions of the pillars produced in PDMS mea sured within 10% of th e pillars of the mold master. Whereas, pillars produced in agarose and gelatin differed from the mold master by more than 10%. This indicates that from a pure pattern replication perspective, a replica leaf made from PDMS would have m ore representative topography of a living leaf . When comparing the replica fidelity of PDMS t o gelatin or agarose. 14 Table 1: Pattern Resolution Measurements . Height and width measurement s were taken from 20 pillars (or wells, as was the case from the PDMS template) . All data is represented a s mean ± standard deviation. Standard deviation has been used to show the scattering in measured values. See Supplementary Information S2 for AFM and optical profilometer image comparison for the PDMS patterned replicati on process. AFM Optical Profilometer ADEX05 Mold Master ADEX05 Mold Master PDMS Template Patterned Agarose Patterned PDMS Patterned Gelatin Height (µm) 4.44 ± 0.16 4.50 ± 0.03 4.18 ± 0.01 2.96 ± 0.60 4.71± 0.01 3.89 ± 0.06 Width (µm) 14.64 ± 0.47 14.94 ± 0.21 15.11 ± 0.52 10.01 ± 0.46 14.00 ± 0.38 13.50 ± 0.48 Fig. 2. Resolution images . (a) Comparison of the photoresist mold master (ADEX05) undertaken by ( i) AFM and an ( ii) optical profilometer. (b) PDMS 10:1 template 3D optical profile. (c) Op tical profiles for ( i) agarose 5 % w/v, (ii) PDMS 10:1 w/w, and ( iii) gelatin 12.5 % w/v. All optical profilometer images were filtered using a 3-point level function in Profilm 3D Software (Filmetrics) . See Supplementary Information S 2 for AFM and optical profilometer image comparison for the patterned PDMS replication process. 15 3.2 Degradation Measurements. Microbiology experiments can take anywhere from a few minutes or hours to days and even weeks. Consequently, a replica material should not degrade duri ng the duration of the experiment as this will result in a moisture and topography changes, which influence s microorganism behaviour .6, 13 Thus, topography changes may prevent certain experiments from being undertake n. For example studies investigating the influence of chemicals or air quality on microorganism behaviour .4 To study material stability , we examined the degradation of the pattern materials ove r three days under two conditions at 30 C (Fig. 3). A temperature of 30 C was selected as it is a common temperature to cultivate environmental bacteria. The two humidity conditions are as follows: (1) 25% relative humidity. This humidity was selected to mimic the humidity often found in dry conditions . (2) 75% relative humidity. This humidity was selected to slow the degradation of the patterned samples . These two relative humidities can be found in cities around the world .58 For patterned agarose a relative humidity of 75% effectively slowed the degradation to the equilibrium point by 36 hours (Fig. 3a). The equilibrium point is defined as the time in which the material reaches an equilibrium moisture exchange with the surrounding environment. Regardless of the concentration of agarose , the equilibrium point (88 ± 1 % weight loss) was reached within 24 h and 48 h for 25% and 75% relative humidity , respectively . For a humidity of 75% no significant difference in the degradation characteristics were observed . Thus, indicating that there was no advantage in increasing or decreasing the concentration of aga rose from 2.5 to 5 % w/v – the concentration of agarose routinely used in microfabricati on.31, 35 In contrast , for gelatin a relative humidity of 75% lowered the equilibrium point (Fig. 3c). Furthermore, r egardless of the concentration of gelatin the equil ibrium point (80 ± 1 % weight loss) was reached within 24 h in both humidity conditions . Similar to agarose , changing the concentration of gelatin in PBS provided no advantage in degradation characteristics. For patterned PDMS no degradation was observed over the 72 h experimental duration in either humidity conditions . In addition , no concentration dependence was observed (Fig. 3b). In summary, the degradation results indicate that a replica leaf produced from PDMS , would be stable in environmental humidi ties that plants occupy around the world. Whereas, agarose and gelatin degrade within 48 h of fabrication in environmental humidity ranges. In literature, agarose and gelatin has been reportedly stored i n water or in a fridge to minimise degradation , respe ctively.44, 46 However, these conditions are not compatible with phyllosphere microbiology experiments , and affect the resolution of microstructures. For instance, storing agarose in water results in swelling and he nce changes the microstructure topography . Furthermore, the majority of microorganisms that reside on plants generally do now grow in low temperatures, such as those experienced in a fridge – 0 to 4 °C. PDMS additionally offers the ability to easily send sample s to laboratories on the other side of the 16 world , as the PDMS replica would not degrade due to the environmental conditions during transportation. Whereas, agarose and gelatin would quickly end up in a liquefied state, unless stored in the aforementio ned unfavourable conditions. Fig. 3. Degradation measurements for the three selected materials. The weight l oss over 72 h for (a) agarose, (b) PDMS , and (c ) gelatin was investigated at relative humidities (rH) of (i) 25% and (ii) 75% at 30 C. Five samp les were measured for each material at each concentration. 17 3.3 Contact Angle Measurements Surface energy and h ydrophobicity in particular, constitute an important physical property of the cuticle of a leaf. A surface is classified as either hydrophilic, hydro phobic , or superhydrophobic, when the contact angle of water is < 90°, > 90°, and >150°, re spectively. We selected enzymatically isolated leaf cuticles from Citrus × aurantium (bitter orange) and Populus × canescens (poplar) plant species to compare our ma terials against (Fig. 4).23 For each of the three replica materials we compared both flat and patterne d surfaces (see Supplementary Fig. S3, for contact angles at different material concentrations) . The hydrophobicity of a surface is important , as the presence of water on a surface impacts resource availability and colonisation patterns of microorganisms in the phyllosphere . Furthermore, the hydrophobicity of the surfaces also influences the microorganism attachment processes. Microorganism s can achieve attachment by (1) adapting to enable attachment , or (2) by forming biofilms 13, 59. Our results indicate that patterned PDMS (95.5 ± 0.6°, N = 5 ) is hydrophobic . Whilst, agarose (15.9 ± 0.9°, N = 5 ) and gelatin (78.3 ± 1.0°, N = 5 ) are hydrophilic . Furthermore, our results are in agreeance with literature .41, 60 Patterning the different materials with our test pattern did not influence the hydrophobic properties of the materials (Fig. 4). This is attributed to the size of the pillars , as there is no change in pinning of the water droplet observed on this scale.61 In addition, changing the concentration also provided no significant ch ange in the hydrophobic ity of each of the materials (Supplementary Fig. S3). Both bitter orange and poplar leaf cuticles were determined to be hydrophobic , with contact angles measured to be 97.9 ± 2.7° (N = 5) and 103.8 ± 0.7° (N = 5), respectively. In ge neral, c ontact angles of plant leaves can vary considerably from hydrophilic to superhydrophobic .32, 62, 63 No significant difference was measured between bitter orange and PDMS . However, a slight difference was measured between Poplar and PDMS (P < 0.05, N = 5) . This is in contrast to the significant difference observed between the leaf cuticles and both agarose and gelatin (Fig. 4). In summary, the hydrophobicity of PDMS is comparable to bitter orange and poplar. Whereas , agarose and gelatin are considerably more hydrophilic. Conversely, PDMS as a replica surface would be a suitable artificial surface for attachment studies of hydrophobic leaves. In addition, the degree of hydrophobicity of PDMS can be modified tem porarily through oxygen plasma or extended in duration with polyvinylpyrrolidone (PVP) treatment . Both modifications are not harmful to microorganisms .64 The use of PVP treatment would enabl e more extensive attachment studies to be undertaken using a PDMS replica surface . An area that has been hig hlighted as requiring more extensive studies to be carried out .13 18 Fig. 4. Contact angle comparison of leaf cuticles with the three selected replica materials. Contact angles of bitter orange and p oplar leaf cuticles , are compared with contact angles of agarose 5 % w/v , PDMS 10:1 w/w and gelatin 12.5 % w/v. See Supplementary Fig. S3 for contact angle measurements for the three concentrations of each material. Data is presented as the mean ± SEM, *P<0.05, **P<0.01, ***P<0.001 , and ****P<0.0001. N.S. indicates no significant difference between measurements. 19 3.4 Bacterial Survival. To assess the suitability of the materials to support microorganism life we used bacterial survival expe riments , with Pantoea agglomerans 299R as our model microorganism (Fig. 5). The bacterium P. agglomerans 299 was isolated from a Bartlett pear tree leaf. Strain P. agglomerans 299R is a spontaneous rifampicin resistant mutant of P. agglomerans 299.52 We selected P. agglomerans 299R as our model microorganism as it is: (1) a model microorganism for leaf colonisation , (2) well characterised and fully sequenced , and (3) it is genetically amendable (able to produce mutants and bioreporters) .27, 52 We compared the bacterial survival characteristics from the replica materials against isolated Citrus × aurantium (bitter orange) leaf cuticles . Thi s was done to determine which material resembled bacterial survival characteristics observed on the surface of lea ves – in the absence of the nutrients from the leaf . This is of importance, as we are looking for a suitable material to form a replica leaf platform for phyllosphere microbiology studies. In the case of agarose , the population of P. agglomerans 299R stabilise d at a normalised cell density of 1.06 ± 0.03 (N = 5) after 24 h (Fig. 5) - regardless of whether the material was patterned or flat (Supp lementary Fig . S4). We conjectured that the population stabilised due to the presence of moisture on the surface of the agarose . As bacterial life can be sustained when moisture is present in the environment . In this case the moisture on the surface of the agarose is a result of the agarose degrading . Furthermore, agarose is derived from red seaweed which is comprised of polysaccharides. Hence, agarose contains a potential nutrient source of carbon for bacteria. However, due to the bacteria not hav ing the n ecessary enzymes to hydrolyse the α-(1 → 3) and β -(1 → 4) glyosidic bonds between the material monomers, this does not provide a nutrient source for bacteria multiplication .65 As a result, no increase in the population of P. agglomerans 299R was observed. Conversely, on gelatin an increase in the population of P. agglomerans 299R was observed in the first 12 h after inoculation ( Fig. 5). This growth wa s attributed to gelatin providing a nutrient source , in the form of pe ptides and proteins. As the bacteria l enzymes were able to hydrolyse the peptide bonds . Thus, producing a suitable nutrient source of carbon and nitrogen. After 12 h the population stabilise d at a normalised cell density of 1.69 ± 0.03 (N = 5). Indicating that this is the largest population density gelatin can sustain. It is important to note that enzymatic break down of gelatin as a nutrient source by the bacteria would also influence the degradation characteristics of the material .66 As this breakdown often results in gelatin becoming liquefied .43 In contrast, P. agglomerans 299R populations were not sustained on either isolated bitter orange cuticle s or PDMS. Once a cuticle is isolated from a leaf, there is no nutrient support from the leaf and the cuticle itself does not provide any nutrients for the P. agglomerans 299R . Hence, no increase in the population of P. agglomerans 299R was observed. Furthermore, with the absence of moisture on the surface of the bitter orange cuticle s, the P. agglomerans 299R died within 6 h (Fig. 5). In the case 20 of PDMS, bacteria do not have the enzymes necessary to degrade the PDMS to form a sustainable nutrient source. PDMS also did not degrade in the experimental conditions ( Fig. 3), indicating th at an exchange of moisture with the environment did not occur . As a result PDMS does not provide the necessary moisture to sustain bacterial life . The lack of nutrients and moisture result ed in the population of P. agglomerans 299R dying within 6 h of inoc ulation to the patterned PDMS surface . Which is comparable to the behaviour observed on the isolated leaf cuticles. In summary, agarose and gelatin will provide a moisture source to sustain bacterial life. In addition, gelatin is able to be degraded by the bacteria to produce a nutrient source that promotes an increase in population , until the maximum population sustainable by gelatin is reached. Thus, indicating that the inherent properties of the material influences bacterial viability . Which also indicat es that the choice of replica material has the p otential to influence phyllosphere microbiology studies. For instance, if one was to examine the influence of nutrient supply, the background supply due to the replica surface material will inter fere with mea surements . Thus , rendering any results obtained unreliable. Conversely, bacterial life was not sustained on either PDMS or bitter orange cuticles . Indicating that PDMS is more representative of a living leaf surface . In addition, modifying PDMS could allow a controlled nutrient or moisture supply to be introduced . For example, his could be achieved with the use of fillers , such as carbon nanotubes. Hence this would enable a nutrient supply more representative of the living leaf while retaining the physical advantages of PDMS .67 21 Fig. 5. Normalised cell density of Pantoea agglomerans 299R. The normalised cell density of P. agglomerans 299R was analysed over 72 h on: Citrus × aurantium (bitter orange) cuticles , agarose 5%, PDMS 10:1 w/w, and gelatin 12.5%. See Supplementary Figure S4 for contact normalised cell density measurements for the three concentrations of each material. Data here is presented as the mean ± SEM . 22 4. Conclusions Our work has demonstrated the potential of PDMS as a replica leaf material for phyl losphere microbiology . At the same time , our results highlighted the drawbacks of agarose and gelatin through comparing o ptical resolution, degradation characteristics, hydrophobic properties, and bacterial survival to PDMS . Using optical profil ometry, we demonstrated that agarose and gelatin replicas would not provide topography comparable to that of a living leaf. Dimensions of the test pillars reproduced in agarose and gelatin were found to differ from the mold master by more than 10%. Furthermore, agaro se and gelatin degraded considerably within 72 h in both high and low humidity conditions at 30 C. On the other hand, the height and width of pillars reproduced in PDMS were within 10% of the dimensions measured for the mold ma ster. Furthermore, PDMS displayed no structural degradation within 72 h. These results indicate that in terms of topography , a replica leaf made from PDMS would be more representative of a leaf surface. In addition, PDMS would not degrade throughout the duration of an experiment . Therefore , will not influenc e microorganism behaviour. The suitability of PDMS was further supported by measured contact angles . The measured contact angles were comparable to those of isolated Citrus × aurantium and Populus × canescens leaf cuticles. In add ition, we examined the influence of the replica materials on bacteria l survival in comparison with isolated Citrus × aurantium leaf cuticle s. These results highlighted that PDMS is representative of a leaf cuticle, as both are unable to support bacterial l ife. Whereas, agarose and gelatin were able to sustain bacterial life . Thu s, indicat ing that these materials would provide a nutrient source . As a result, render ing these materials unsuitable for nutrient supply studies. In summary , the results presented here indicat e that in contrast to agarose and gelatin, the combined properties of PDMS make for a suitable replica material for phyllosphere microbiology . Our experiments highlight the importance of considering the inherent material properties when selecti ng a material as a replica surface. In our current work , we are investigating the preparation of biomimetic leaf replicas made from PDMS with tailored nutrient permeability for plant -microbe interactions at a single -cell resolution. Acknowledgements This work was funded by the Biomolecular Interaction Centre and Marsden Grant UOC1704. R.S. and N.A. thank the National Science Challenge - Science for Technological Innovation for Post -Doctoral Fellowship and Doctoral Scholarship funding. We thank Helen Dever eux and Gary Turner for technical assistance. 23