Priscilla B. Dourado II ; Maria C. Quecine II ; Fernando D. Azevedo II ; Aline A. Pizzirani-Kleiner II. Two endophytic strains of Methylobacterium spp. Results show that biofilm formation is variable and that plant surface and possibly root exudates have a role in Methylobacterium spp.
Key words: Methylobacterium, endophytes, biofilm; sugarcane. Members of this genus degrade one-carbon compounds C1 such as methanol and methylamine 8 and are widely distributed in the environment, colonizing soil, water, leaf surfaces, nodules, seeds, air, and sediments 9. They have been shown to interact symbiotically with different plant species of agronomic importance 2, 3, 5. Specifically in sugarcane, Methylobacterium spp. Better understanding of Methylobacterium spp. The aim of this study was to investigate the role of the living plant surface in biofilm formation by M.
The strains SR1. They were previously isolated from citrus and characterized 3. Bacterial fresh culture were harvested by centrifugation 10 min at xg and the pellet subsequently resuspended and diluted in PBS phosphate-buffered saline, pH7.
Commercial wooden sticks were used with the same conditions as a control. Three repetitions of each treatment were used. Andreote et al. Here we have investigated whether the strains might have different capacities for forming biofilms on different surfaces, thus possibly determining their ability to become endophytic colonizers of sugarcane plants. Our observations of wooden sticks showed biofilm formation only by M. Cells of M. Biofilm formation by the strain AR1.
It is remarkable that, even after a short period of incubation 3 DAI the treatment with the strain AR1. The formation of biofilm on an inert material such as the wooden stick used here shows that biofilm formation of strain AR1. On the sugarcane seedling roots, biofilm formation by both strains AR1. In contrast to what was observed for wooden sticks, strain SR1.
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In the control treatment without bacterial cells inoculation of all experiments no bacteria were observed by similar material preparation and SEM scanning data not shown. Quantification of the Methylobacterium spp. For quantification, roots of the plants were submerged in 2 mL of PBS and agitate at rpm for 1 hour with the shoots remaining above the buffer. The resulting cell suspension was diluted and plated to quantify the bacteria in CHOI 3. The disinfected material was separated in root and shoot, weighed and grinded in 2 mL of PBS buffer.
The microbial transglutaminase mTG, Ajinomoto Co. Then 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 on the PDMS template and left at room temperature for five minutes to stabilise. The patterned gelatin was then carefully peeled off the PDMS template [ 44 ]. All optical profiles of the patterned material samples 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. For agarose and gelatin the high sidewalls of the pillars made imaging difficult due to light scattering. To correct for this, the data was processed using the inbuilt remove outliers function in the Profilm3D software ver.
For the remove outliers function, the invalid pixels filled in method was used and the maximum slope was set to During post processing, all images were filtered using a 3-point level function in Profilm3D software. This was done to account for any non-level placement of the samples on the Profilm 3D stage. AFM images were not possible for agarose and gelatin due to the patterned agarose and gelatin degrading during the required imagining period, and the AFM tip losing contact.
The humidity levels were selected to: i mimic dry conditions similar to a climate-controlled laboratory; and ii 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 concentrations for each material were used. Each sample had an initial diameter of Weight measurements were taken at 0, 2, 4, 6, 12, 24, 48, and 72 hours.
A weightless percentage measure was then determined for each of the time points. 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. This environment must be favourable for microorganisms, and minimise any potential swelling of the materials. To achieve this, a saturated salt solution was prepared by dissolving 72 g of sodium chloride , Sigma-Aldrich in mL of deionised water.
The saturated salt solution was then placed next to the patterned material samples in a 7 L airtight container. Flat samples were compared against patterned samples for each material at the three selected concentrations. Five samples with a diameter of This diameter was selected to minimise the potential occurrence of edge effects on the water droplets. Prior to the contact angle measurements, all samples were dried with dry nitrogen gas to minimise any potential effects from surface moisture.
In addition, experiments were conducted within an hour of peeling the material off the PDMS template. The surface energy of all the materials was determined using deionised water. P values less than 0. The P. The supernatant was discarded and the bacteria was suspended in fresh phosphate buffer to an OD nm of 0. Five samples were taken per time point and all samples were sterilised for 15 minutes with ultra-violet sterilisation. Five samples were randomly selected per measurement time points corresponding to 0, 2, 4, 6, 12, 24, 48, and 72 hours. Each individual sample was suspended in 1 mL of fresh phosphate buffer in a 1.
The samples were left to cool for five minutes to minimise any potential thermal stress and then vortexed for another five minutes. The recovered supernatant was then plated onto nutrient agar by undertaking serial dilutions using PBS. Colony-forming units CFU were expressed as bacterial cell density per unit area for each sample 7. The cell density was normalised to the initial number of cells after inoculation.
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This was undertaken to enable a comparison between the survival curves of the bacterial colonies on the different materials. The microstructures found on leaves are dependent on the plant species and can range in complexity. Microstructures found on leaves include groves and stomata pores to the more complex structure of trichomes leaf hairs. The ability to sufficiently replicate the surface topography of a living leaf is important for producing replica leaf surfaces that can be used as a control surface in phyllosphere microbiology. This is of significance, as the topography of the leaf surface has the ability to influence colonisation and attachment behaviours of microorganisms [ 4 , 20 ].
To investigate the capability of our selected materials to replicate the complex topography of leaves, we used a test pattern. Such dimensions are comparable to the features found on leaf surfaces—trichome, stomata, and grooves. Resolution comparisons of the patterned materials were undertaken using a 3D optical profilometer Fig 2.
Optical profiles were selected over more commonly used techniques to compare microstructure resolution, such as AFM and scanning electron microscopy SEM. The use of these techniques was hindered by the softness and quick degradation of agarose and gelatin. When investigating the suitability of AFM imaging, we regularly observed the AFM tip losing contact with the sample during imaging.
The use of SEM imaging was limited by the requirement of a conductive coating, as both agarose and gelatin are non-conductive. To produce a conductive coating, the samples would have to be placed under vacuum. However, tests with a short three-minute vacuum resulted in a noticeable structural degradation in both materials. Conversely, obtaining a 3D optical profile requires no modifications to be made to the materials, and images can be acquired in approximately three minutes; thus, minimising any degradation affects see section 2.
All optical profilometer images were filtered using a 3-point level function in Profilm3D Software Filmetrics.
See S2 Fig. For example, the height of the mold master was measured to be 4. Furthermore, the measured widths were In consequence, this indicates that no significant difference existed between 3D profiles taken by either AFM or an optical profilometer. Height and width measurements were taken from 20 pillars or wells, as was the case from the PDMS template.
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Standard deviation has been used to show the scattering in measured values. A summary of the 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.
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 smaller. The height and width of the pillars in gelatin were 0. We conjectured that this was due to the gelatin degrading while setting. 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, and placed the gelatin and PDMS template in a fridge prior to peeling the gelatin off [ 44 ]. Furthermore, from our experience the trichomes leaf hairs would not withstand being peeled off the PDMS template even following this protocol, as they are inherently fragile [ 20 ]. Conversely, the pillars replicated into PDMs were more representative of the mold master. This indicates, that from a pure pattern replication perspective, a replica leaf made from PDMS would be more representative of the living leaf topography, than a replica made from gelatin or agarose.
Microbiology experiments can take anywhere from a few minutes or hours to days and even weeks. Topography changes, in particular, may prevent certain experiments from being undertaken, such as studies investigating the influence of chemicals or air quality [ 4 ]. Furthermore, these two relative humidities can be found in cities around the world [ 58 ]. Five samples were measured for each material at each concentration. We defined the equilibrium point as the time in which the material reaches an equilibrium moisture exchange with the surrounding environment.
This indicated that there is no advantage in changing the concentration of agarose—between 2. Similar to agarose, changing the concentration of gelatin in PBS did not improve the degradation characteristics. For patterned PDMS no degradation was observed over the 72 hours experimental duration in either humidity conditions and 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 humidities that plants occupy around the world.
Whereas, agarose and gelatin degrade within 48 hours of fabrication in environmental humidity ranges. In literature, agarose and gelatin have been stored in water or in a fridge to minimise degradation, respectively [ 44 , 46 ]. However, these conditions are not compatible with phyllosphere microbiology experiments and also affect the resolution of the replicated microstructures. For instance, storing agarose in water results in swelling and hence changes the microstructure topography.
On the other hand, replica leaves produced in PDMS would allow samples to be sent to laboratories on the other side of the world. As PDMS replica leaves would not degrade in the typical environmental conditions experienced during transportation. Unless stored in the aforementioned unfavourable conditions, transportation would quickly liquefy agarose and gelatin replica leaves.
The hydrophobicity of a leaf surface is another important property that needs to be considered when selecting a suitable replica material.
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As the presence of water on a leaf surface impacts resource availability and colonisation patterns of microorganisms in the phyllosphere. Furthermore, the hydrophobicity of the leaf surface influences the microorganism attachment processes. Microorganisms can achieve surface attachment by adapting to enable attachment or by forming biofilms [ 13 , 59 ]. In addition, we compared both flat and patterned surfaces for each of the three replica materials see S3 Fig , for contact angles at different material concentrations.
See S3 Fig for contact angle measurements for the three concentrations of each material. Our results indicated that the patterned PDMS These results are in agreeance with literature [ 41 , 60 ]. Furthermore patterning the different materials with our test pattern did not influence the hydrophobic properties of the materials Fig 4. This was attributed to the size of the pillars, as there is no change in the pinning of the water droplets on this scale [ 61 ]. In addition, changing the concentration of the materials did not change the hydrophobicity of the materials S3 Fig.
For comparison, isolated bitter orange and poplar leaf cuticles used, and they were determined to be hydrophobic, with contact angles measured to be In general, contact angles of plant leaves can vary considerably from hydrophilic to superhydrophobic [ 32 , 62 , 63 ]. In our case, no significant difference in contact angle was measured between bitter orange and PDMS. Nonetheless, these results provide a significant contrast to the difference observed between the leaf cuticles and both agarose and gelatin Fig 4. In summary, the hydrophobicity of PDMS was comparable to bitter orange and poplar.
Whereas, agarose and gelatin were considerably more hydrophilic. Thus indicating PDMS is a suitable replica surface material for conducting attachment studies of hydrophobic leaves.
In addition, the degree of hydrophobicity of PDMS can be modified temporarily through oxygen plasma or extended in duration with polyvinylpyrrolidone PVP treatment. Both modifications are not harmful to microorganisms [ 64 ]. In literature this has been highlight as an area that requires more extensive studies to be undertaken [ 13 ]. To assess the suitability of the materials to support microorganism life we conducted bacterial survival experiments, with Pantoea agglomerans R as our model microorganism Fig 5. The bacterium P.
Strain P. We selected P. We used isolated leaf cuticles, as we wanted to determine which material closely resembled the bacterial survival characteristics observed on the surface of leaves, in the absence of the nutrients supplied from the leaf. This was important, as we were looking for a suitable material to form a replica leaf platform for phyllosphere microbiology studies. The normalised cell density of P. See S4 Fig for contact normalised cell density measurements for the three concentrations of each material. In the case of agarose, the population of P.
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 was a result of the agarose degrading. Furthermore, agarose is a potential source of nutrients for bacteria, due to agarose being derived from red seaweed which is comprised of polysaccharides. However, P.
Thus, we conjectured that the population of P. Conversely, on gelatin an increase in the population of P. This growth was attributed to gelatin providing a nutrient source in the form of peptides and proteins. As the bacterial enzymes were able to hydrolyse the peptide bonds, gelatin provided a suitable nutrient source of carbon and nitrogen. After 12 hours the population stabilised at a normalised cell density of 1. Thus, indicating that this is the largest population density gelatin can sustain. It is important to note that the enzymatic breakdown of gelatin as a nutrient source by the bacteria would influence the degradation characteristics of the material [ 66 ].
After all, this breakdown often results in gelatin becoming liquefied [ 43 ]. In contrast, P. 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. Hence, no increase in the population of P.
Granule formation in early infection
Furthermore, with the absence of moisture on the surface of the bitter orange cuticles, the P. PDMS also did not degrade in the experimental conditions Fig 3 , indicating that an exchange of moisture with the environment did not occur. As a result, PDMS did not provide the necessary moisture to sustain bacterial life. The lack of nutrients and moisture resulted in the population of P.
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 can be degraded by P. Thus, indicating that the inherent properties of agarose and gelatin influences bacterial viability. Which also influences the choice of replica leaf material that can be used for phyllosphere microbiology studies. For instance, any inherent nutrient source from the replica material would influence nutrient supply studies.
Hence any results obtained from such a system would be deemed unreliable. Conversely, bacterial life was not sustained on either PDMS or bitter orange cuticles. This indicated that PDMS is more representative of a living leaf surface. In addition, through suitable modification PDMS could enable a controlled nutrient or moisture supply to be introduced.
For example, his could be achieved with the use of fillers such as carbon nanotubes. This would enable a nutrient supply more representative of the living leaf, while retaining the physical advantages of PDMS [ 67 ]. Our work has demonstrated the potential of PDMS as a replica leaf material for phyllosphere microbiology. At the same time, our results highlighted the drawbacks of agarose and gelatin through comparing optical resolution, degradation characteristics, hydrophobic properties and bacterial survival to PDMS.
Using optical profilometry, we demonstrated that agarose and gelatin replicas would not provide topography comparable to that of a living leaf. PDMS also displayed no structural degradation within 72 hours. These results indicate that in terms of topography, a replica leaf made from PDMS would be more representative of a leaf surface. On the other hand agarose and gelatin were both able to sustain bacterial life, thus indicating that these materials would influence microorganism behaviour. As a result, both of these materials must be considered unsuitable for nutrient supply studies.
In summary, the results presented here indicate 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 selecting 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. Bacteria viability for bacteria at i different concentrations, for a agarose, b PDMS, and c gelatin. Lukas Schreiber University of Bonn, Germany for the kind gift of the isolated cuticles used in this study. Browse Subject Areas? Click through the PLOS taxonomy to find articles in your field. Abstract 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.
Download: PPT. Fig 3. Degradation measurements for the three selected materials. Fig 4. Contact angle comparison of leaf cuticles with the three selected replica materials. Fig 5. Normalised cell density of Pantoea agglomerans R. Supporting information. S1 Fig.