spatial distribution analyses of natural phyllosphere-colonizing bacteria on a rabidopsis thaliana...

Remus-Emsermann et al.

This article is protected by copyright. All rights reserved.

Spatial distribution analyses of natural phyllosphere-colonizing bacteria 1

on Arabidopsis thaliana revealed by fluorescence in situ hybridization1 2

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Mitja N.P. Remus-Emsermann1*, Sebastian Lücker2, Daniel B. Müller1, Eva 5

Potthoff1, Holger Daims2, and Julia A. Vorholt1* 6

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1Institute of Microbiology, ETH Zurich, Switzerland 8

2Division of Microbial Ecology, Department of Microbiology and Ecosystem 9

Science, University of Vienna, Altanstrasse 14, 1090 Vienna, Austria 10

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*Corresponding authors 12

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Running title: Bacterial distribution on Arabidopsis phylloplanes 14

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Subject Category: Microbial population and community ecology 16

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Keywords: environmental heterogeneity/ fluorescence in situ hybridization/ 23

phyllosphere/ single-cell microbiology/ community composition 24

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This article has been accepted for publication and undergone full peer review but has not been through the

copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version

and the Version of Record. Please cite this article as doi: 10.1111/1462-2920.12482

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26 Abstract 27

Bacterial colonizers of the aerial parts of plants, or phyllosphere, have been 28

identified on a number of different plants using cultivation-dependent and 29

independent methods. However, the spatial distribution at the micrometer scale 30

of different main phylogenetic lineages is not well documented and mostly based 31

on fluorescence-tagged model strains. In this study we developed and applied a 32

spatial explicit approach that allowed the use of fluorescence in situ 33

hybridization (FISH) to study bacterial phylloplane communities of 34

environmentally grown Arabidopsis thaliana. We found on average 5.4×106 35

bacteria per cm2 leaf surface and 1.5×108 bacteria per gram fresh weight. 36

Furthermore we found that the total biomass in the phylloplane was normally 37

distributed. About 31% of the bacteria found in the phylloplane did not hybridize 38

to FISH probes but exhibited infrared autofluorescence indicative for aerobic 39

anoxygenic phototrophs. Four sets of FISH probes targeting Alphaproteobacteria, 40

Betaproteobacteria, Actinobacteria, and Bacteroidetes were sufficient to identify 41

all other major contributors of the phylloplane community based on general 42

bacterial probing. Spatial aggregation patterns were observed for all probe-43

targeted populations at distances up to 7 µm, with stronger tendencies to co-44

aggregate for members of the same phylogenetic group. Our findings contribute 45

to a bottom-up description of leaf surface community composition. 46

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48 Introduction 49

The above ground organs of plants, or phyllosphere, represent a large global 50

habitat for microorganisms. The habitat is dominated by leaves, which were 51

estimated to encompass as much as 109 km2 world-wide and the total number of 52

inhabitants in the phyllosphere is believed to be as high as 1026 bacteria (Morris 53

and Kinkel, 2002). Microorganisms that are living on the surface of leaves, also 54

known as the phylloplane, are generally referred to as epiphytes. Many of these 55

epiphytes are not harmful to the host plants (Vorholt, 2012), but instead are 56

commensals or are even beneficial to the plant host by preventing the growth of 57

secondary colonizers that might be pathogens (Stockwell et al., 2010; Innerebner 58

et al., 2011). 59

Cultivation-dependent and more recently independent methods were used to 60

describe natural community compositions on leaves of various plants (Yang et 61

al., 2001; Delmotte et al., 2009; Knief et al., 2010b; Knief et al., 2012); for reviews 62

see Vorholt (2012); Rastogi et al. (2013), and for the model plant Arabidopsis 63

thaliana in particular see Delmotte et al. (2009); Vorholt (2012); Bodenhausen et 64

al. (2013). Next generation sequencing approaches showed that the bacterial 65

communities found on different plant species are plant-host dependent, stable 66

and have a high degree of reproducibility at the genus level or higher (Knief et al., 67

2010b; Redford et al., 2010; Vorholt, 2012). These findings suggest that the plant 68

host and colonizers might select for each other. However, the underlying 69

processes of community assembly remain to be elucidated. 70

Current knowledge of bacterial colonization of the phylloplane at a micrometer 71

or single-cell scale is mostly based on studies that investigated phylloplane 72 Acc

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colonization using fluorescently tagged model strains (Leveau and Lindow, 2001; 73

Brandl and Mandrell, 2002; Monier and Lindow, 2004, 2005b, a; Zhang et al., 74

2009; Tecon and Leveau, 2012). From these and other studies it became 75

apparent that the phylloplane is not uniformly colonized, but that bacterial 76

colonization takes place in hotspots (Kinkel et al., 1995; Monier and Lindow, 77

2004, 2005b; Sy et al., 2005; Remus-Emsermann et al., 2012). Environmental 78

studies using general DNA dyes and scanning electron microscopy, as well as 79

fluorescent bioreporter-based studies, have shown that bacteria prevalently 80

colonize trichomes, stomates, hydatodes and other prominent leaf features as 81

well as grooves formed by epidermis cells (Beattie and Lindow, 1999; Lindow 82

and Brandl, 2003; Monier and Lindow, 2004). To date, studies investigating 83

natural phylloplane communities at a micrometer scale have been scarce (Morris 84

et al., 1997; Fett and Cooke, 2003; Warner et al., 2008), and efforts to 85

taxonomically identify individual epiphytes at a micrometer scale have been 86

made even less frequently (Li et al., 1997; Bisha and Brehm-Stecher, 2009). In 87

particular, no information on natural epiphytic communities has been obtained 88

by using a multi-labeled combinatorial FISH approach. In this study, we aimed at 89

analyzing natural communities on leaves of Arabidopsis thaliana, which is the 90

best-studied plant model system and one of the current models for microbiota 91

colonization both above- and below ground (Innerebner et al., 2011; Bulgarelli et 92

al., 2012; Lundberg et al., 2012; Vogel et al., 2012). We set out to address the 93

following issues: Firstly, we wanted to understand whether and how the overall 94

bacterial community composition which was described at a low spatial 95

resolution, i.e. the level of individual plants (Knief et al., 2010b; Redford et al., 96

2010), is reflected at micrometer-scale resolution; secondly, if phylloplane 97 Acc

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colonization patterns are predictable, e.g., if the different phylloplane colonizers 98

have non-random spatial relationships, as it was shown for dual species 99

phyllosphere colonization by Monier and Lindow (2005b); and thirdly, if the 100

preferential colonization of specific leaf organs as observed for single-strain 101

inoculations can also be seen in bacterial communities of environmentally grown 102

Arabidopsis. 103

To achieve these goals we tested several methods and established a protocol that 104

allowed us to investigate the distribution of bacterial groups on leaf surfaces at a 105

micrometer scale without introducing a spatial bias. Bacteria were recovered 106

from plant leaves using adhesive tape before fluorescence in situ hybridization 107

(FISH) was used to visualize the distribution of the phylloplane community at the 108

single-cell level. Analysis of the obtained patterns was performed using 109

computer assisted image analysis, and spatial statistical tools provided a 110

systematic microscale investigation of representative parts of the phylloplane of 111

environmentally grown, naturally colonized Arabidopsis leaves. 112

113

Materials and methods: 114

Sampling of environmentally grown Arabidopsis thaliana and isolation of 115

the epiphytic community by cuticle tape lift 116

Arabidopsis plants from a wild population were sampled by digging out whole 117

plants including roots and soil (July 2013, Windisch AG, Switzerland; GPS 118

coordinates 47° 28.898’, 8° 13.054’). Plants were transported to the lab on wet 119

tissue. Weather conditions were hot and dry during sampling, i.e. the minimal 120

night temperature was 15 °C and the maximal day temperatures 32 °C. 121 Acc

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Cuticle tape lifts were performed while the plants were still turgescent. To this 122

end, double-sided adhesive tapes (Yamori 50, 3M double sided adhesive tape 123

9086, X-Film Montex DXP 12, Tesa double sided adhesive tape, two Neschen 124

double sided tape “strong/strong”, and “strong/removable”, all tapes were 125

acquired at Modulor, Berlin, Germany) were glued onto microscopical slides with 126

the protective sheath still intact on the backside of the tape. The protective 127

sheath of the tape was removed and the leaf was carefully flattened onto the 128

sticky layer of the tape using an ethanol wiped, clean glass stick with 1 mm 129

diameter. Spring steel tweezers were used to rip off the leaf from the sticky tape, 130

which resulted consistently in an imprint of the leaf on the adhesive tape, which 131

was checked by phase contrast microscopy. If the leaf remained intact, its still 132

untreated side was then applied to another adhesive tape, which resulted in 133

combined adaxial and abaxial samples of the same leaf. 134

Transferred microorganisms on the adhesive tapes were fixed using a modified 135

protocol of Daims et al. (Daims et al., 2005) by application of 4% 136

paraformaldehyde solution (PFA, 4% PFA in 1 x phosphate buffered saline, pH 7) 137

on top of the adhesive tape harboring the recovered bacteria and incubation for 138

3 hours at room temperature in sealed incubation chambers. After incubation, 139

the slides were dipped for 10 seconds into sterile double distilled H2O to wash 140

off the PFA solution. The slides were then dehydrated by incubations for three 141

minutes each in 50%, 80%, and 100% ethanol. After drying the slides with 142

compressed air and an additional drying for 5 minutes in the dark, the slides 143

were stored in tightly closed plastic tubes at -20 °C in the dark until they were 144

processed for microscopy. 145

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Fluorescence in situ hybridization 147

Ribosomal RNA-targeted oligonucleotide probes designed and evaluated for 148

different phylogenetic groups were chosen from the ProbeBase database (Loy et 149

al., 2007) (for a general discussion on limitations of probes see Amann and Fuchs 150

2008) based on available data for Arabidopsis phyllosphere communities 151

(Delmotte et al., 2009; Knief et al., 2010b; Vorholt, 2012). Probes were 5'-tagged 152

with the fluorochromes Cy3, Cy5, 5’FAM, or ATTO488 (Table 1). Hybridization 153

conditions were chosen according to ProbeBase and the hybridization protocol 154

was adapted from Daims et al. 2005 (Daims et al., 2005). In short, frozen samples 155

were allowed to warm to room temperature in the dark to prevent precipitation 156

of air humidity before they were removed from their plastic containers. Samples 157

were once more dehydrated in an ethanol series as described above to remove 158

any residual water. Remaining ethanol was blown off using compressed air and 159

the slides were allowed to dry in the dark for 10 minutes. Depending on the size 160

of the leaf imprints, the samples were subsequently overlaid by 10 to 40 µL 161

hybridization buffer. Stringent hybridization conditions were ensured by adding 162

formamide to the final concentrations indicated by ProbeBase (see Table 1). All 163

Cy3, Cy5 and ATTO488 labeled probes except HGC69A were added to a final 164

concentration of 1.5 ng/µL, 5’FAM labeled probes to a concentration of 2.5 165

ng/µL. Probe HGC69A was used at a concentration of 2.25 ng/µL. If necessary, 166

unlabeled competitor oligonucleotides were added at the same concentrations as 167

the respective labeled probes. Samples were then transferred to a hybridization 168

chamber filled with tissue paper soaked in hybridization buffer and hybridized at 169

46 °C for three hours. To remove any non-specifically bound probes, samples 170

were then washed for 10 minutes in the appropriate washing buffer (Daims et 171 Acc

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al., 2005). Slides were then dipped into ice-cold water, dried using compressed 172

air and additionally kept for 10 minutes in the dark to allow water residuals to 173

evaporate. In total we hybridized 14 leaves from 4 plants with the general 174

bacterial probe EUB338 and two group-specific probes each. At least 5 175

hybridization experiments per phylogentic group were performed. 176

Even though only the probe EUB338 instead of the probe mix EUB338 I-III 177

(Daims et al., 1999) was used for general bacterial probing, phase contrast 178

microscopy indicated that all microorganisms present on the tape lift (except for 179

infrared autofluorescent bacteria, see below) were detected. 180

181

Microscopy 182

Microscopy was performed using an AxioObserver D2 epifluorescence 183

microscope (Carl Zeiss GmbH, Oberkochen, Germany) with attached light source 184

X-Cite 120Q (Lumen Dynamics Group Inc., Mississauga, Canada). FAM and 185

ATTO488 emission were visualized using Zeiss filterset 38 HE (BP 470/40 186

nm/FT 495 nm/ BP 525/50 nm), Cy3 emission was visualized using Zeiss 187

filterset 43 HE (BP 550/25 nm/FT 570 nm/ BP 605/70 nm), Cy5 emission was 188

visualized using Zeiss filterset 50 (BP 640/30 nm/FT 660 nm/BP 690/50 nm), 189

and emission of infrared fluorescent bacteria was visualized using a custom filter 190

set with a 320-650 nm excitation filter (BG39, Schott AG, Mainz, Germany), 650 191

nm dichroic mirror (5650dcxru, Chroma, Bellows Falls, VT, USA), and a 850 nm 192

longpass emission filter (RG840, Schott AG). Samples were mounted in Citifluor 193

AF1 antifade (Citifluor Ltd., London, UK) and covered with a coverslip prior to 194

observation with a Zeiss 100 x EC Plan-Neofluar oil-objective (1.3 NA, Phaco 3). 195

Images were acquired using an AxioCam Mrm (Carl Zeiss GmbH) and the 196 Acc

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program AxioVision 4.8 (Carl Zeiss GmbH). For every sample, between 20 and 60 197

random fields of view (FOV) with a size of 83 x 62 µm were recorded with up to 5 198

channels assigned to phase contrast, different fluorescent labels, and infrared 199

autofluorescence. 200

201

Image processing and analysis 202

Images were exported from the AxioVision software to 8-bit greyscale TIFF 203

images. Images of green probe signals generally required more processing due to 204

a low signal to noise ratio. These images were processed using the program Fiji 205

(Schindelin et al., 2012) to reduce the background by performing a rolling ball 206

subtraction and applying a median filter before they were imported into the 207

image analysis software DAIME 2.0 (Daims et al., 2006). Images of infrared 208

fluorescent bacteria likely engaged in aerobic anoxygenic photosynthesis were 209

deconvolved using the Huygens remote manager v.2.1 implementation of the 210

Huygens deconvolution software package (Scientific Volume Imaging, Hilversum, 211

The Netherlands) since the focus layer of the used filterset was different from all 212

other images. All other images were directly imported into DAIME 2.0 where a 213

“blur and subtract” background correction was performed and histogram 214

stretching was applied to improve the results of subsequent image segmentation. 215

Images were segmented in DAIME using the automatic threshold algorithms 216

“isodata” or “local” depending on the individual dataset. Automatically 217

segmented images were manually validated. To exclude non-microbial 218

fluorescent particles from further analysis, the phase contrast images of the 219

affected FOVs were used to check dubious fluorescence signals. False negative 220

results (i.e., cells overlooked during the automated segmentation procedure) 221 Acc

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occurred in case of large differences in fluorescence intensity between cells in 222

the same FOV and were included by using manual segmentation tools provided 223

in DAIME. The image area occupied by bacteria was used as a proxy for bacterial 224

population density. 225

Total bacterial population density was estimated by cumulating the areas 226

occupied by bacterial cells hybridized to EUB338 and by bacteria exhibiting 227

infrared autofluorescence, which did not hybridize to FISH probes. 228

The relative abundance per FOV of a bacterial group under study was estimated 229

by quantifying the area fraction relative to all bacterial cells. To estimate 230

absolute cell numbers of each bacterial group per cm2 leaf surface, the median 231

object area of each probe-target group was measured assuming that the cells 232

formed a monolayer. Subsequently, the corresponding total area was divided by 233

the median object area. 234

To quantify the spatial arrangement patterns of the different phyla and classes 235

under study, we performed a stereological analysis using the linear dipole 236

algorithm implemented in DAIME, which yields pair correlation and pair cross-237

correlation plots with 95% confidence intervals (Daims and Wagner, 2011). 238

Spatial arrangement patterns within the same probe-target group were 239

quantified by taking into account also co-aggregation signals within the same 240

objects to appropriately analyze microcolonies of densely packed cells that could 241

not be further resolved by the segmentation algorithms. To mark the length of an 242

individual cell and thus distances from which pair correlations become 243

meaningful, a vertical line was added to the resulting pair correlation plots to 244

depict the determined average length of the bacteria under study. Pair (cross) 245

correlation values that are not significantly different from one indicate that the 246 Acc

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spatial arrangement of the tested groups is random at the respective distance. 247

Values significantly greater than one indicate co-aggregation, whereas values 248

significantly smaller than one indicate a negative spatial correlation. 249

250

Results 251

Establishment of cuticle tape lifts to observe phylloplane microbial 252

communities 253

To analyze spatial relations of phylloplane bacteria we established a protocol, 254

which consists of applying double-sided adhesive tape to Arabidopsis leaves. A 255

similar approach was previously reported by Bisha and Brehm-Stecher for 256

tomato fruits (Bisha and Brehm-Stecher, 2009). The protocol allowed to readily 257

detect the presence of bacteria by phase contrast microscopy and projected the 258

three-dimensional leaf surface onto a two-dimensional plane, thereby also 259

facilitating image acquisition. In total we screened six different adhesive tapes 260

(see Materials and Methods). Cuticle tape lifts to remove phylloplane bacteria 261

were first applied to Arabidopsis plants cultivated in growth chambers as 262

described before (Innerebner et al., 2011) that were inoculated with red-263

fluorescent mCherry-protein expressing Sphingomonas Fr1 (unpublished). All 264

adhesive tapes tested were able to quantitatively remove the phylloplane 265

community from Arabidopsis leaves resulting in leaves that were almost devoid 266

of fluorescent bacteria (exemplarily shown in Supplemental Figure 1). The 267

bacterial patterns that were observed on the tape lifts matched those 268

colonization patterns that were visible during direct in planta observations of the 269

same fluorescent bacteria (Supplemental Figure 1). The tape “Yamori 50” had the 270

best properties for the study. The autofluorescent background of this tape was 271 Acc

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low and even, in contrast to most other tapes that exhibited high background 272

fluorescence or contained a mesh of fibers that produced an uneven background 273

and severely hampered validation by phase contrast microscopy. 274

As alternative approaches we also tested a 3D-FISH protocol where entire 275

samples were embedded in polyacrylamide pads (Daims et al., 2006) and an 276

analogous technique using agar (Brandl et al., 2006). However, these approaches 277

did not yield satisfactory results as they produced obvious artifacts in the 278

distribution of phylloplane bacteria under study, or the protective gel cover 279

dissolved during the required washing and drying steps. Another obstacle of 280

these approaches was strong uneven autofluorescence from the plant tissue. 281

282

Establishment of FISH procedures 283

In a first step, we used pure culture isolates of typical phyllosphere colonizers to 284

test the FISH probe set (see Materials and Methods, Table 1). The tested strains 285

included the alphaproteobacteria Sphingomonas strain FR1 (Innerebner et al., 286

2011), S. phyllosphaere (Rivas et al., 2004), Methylobacterium extorquens PA1 287

(Knief et al., 2010a), Methylobacterium sp. L04, Methylobacterium sp. P01, and 288

Methylobacterium sp. Q12 (Stiefel et al., 2013), the gammaproteobacterium 289

Pseudomonas syrinage pv. tomato DC3000 (Cuppels, 1986), as well as recently 290

isolated strains including the betaproteobacterium Variovorax sp., the 291

actinobacterium Arthrobacter sp., and the Bacteroidetes member Pedobacter A03 292

(laboratory collection, unpublished). All probes were successfully tested for their 293

ability to detect the specific strains and to discriminate against members of non-294

target phylogenetic groups (examples of hybridizations are shown in 295

Supplemental Figure 2 and 3). 296 Acc

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In contrast to all other bacterial strains tested, we found that the 297

Methylobacterium strains exhibited an erratic hybridization pattern where some 298

cells of a population hybridized very well to FISH probes, while others did not 299

hybridize at all (even when two distinct FISH probes were applied that should 300

both target these cells, i.e. EUB338 and ALF698a; Supplemental Figure 3). 301

Notably, these bacterial strains exhibited infrared autofluorescence, which was 302

previously linked to aerobic anoxygenic phototrophs (AAnPs), after growth in 303

planta as well as on R2A media exposed to a diurnal cycle (Stiefel et al., 2013). 304

The cause of the failure to consistently hybridize to FISH probes is currently 305

unclear and it remains to be shown whether membrane compounds such as 306

hopanoids (Knani et al., 1994; Bradley et al., 2010; Muller et al., 2011), or the 307

expressed photosystems found in the membranes of Methylobacteria might 308

affect diffusibility of the probes or if the cellular ribosome content might be 309

strongly reduced in many, but not all, Methylobacterium individuals. At this time, 310

we can also not completely rule out that the bacterial photosystems are 311

quenching the fluorescent signal of the FISH probes. However, in our pure 312

culture hybridization we did not observe an inverse correlation between high 313

infrared autofluorescence and FISH signal. 314

315

General observation of cuticle tape lifts of the phylloplane 316

Upper and lower phylloplane samples of the same leaves were investigated and 317

the general bacterial cell density was estimated microscopically after hybridizing 318

the samples to the general bacterial probe EUB338. Images of random FOVs were 319

acquired with up to 5 color channels, and a total of more than 1100 FOVs were 320

analyzed. We found that on average, approximately 5% of the abaxial 321 Acc

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phylloplane, i.e. the lower leaf surface, were covered with bacteria (Figure 1), 322

while we hardly detected bacterial cells on the adaxial phylloplane. We therefore 323

focused on the abaxial leaf side for stereological analyses. The bacteria on the 324

abaxial phylloplane were organized in clusters and individual cells (Figure 2). In 325

general, most detected bacteria of the same phylogenetic probe-target group 326

exhibited similar morphologies. Three-dimensional assemblages were observed 327

occasionally (e.g., ball-shaped colonies of Bacteroidetes, Figure 2 C), possibly 328

leading to an underestimation of the total bacterial abundance as measured here 329

from areas in two-dimensional images. We found that bacteria regularly 330

accumulated close to imprints of stomatal openings (Figure 2 A and B) and 331

bacteria of the same phylogenetic group often formed small microcolonies of five 332

or more cells (Figure 2). However, it was common to find cells of different 333

phylogenetic groups to co-aggregate. Bacterial densities increased towards the 334

leave edges and high densities of AAnPs could be found here often (data not 335

shown). 336

The area covered by bacteria per FOV was found to be normally distributed 337

(Figure 1). The standard deviation, as indicated by the slope of the plots, was 338

almost identical between samples (Figure 1). 339

Members of the Alphaproteobacteria, Betaproteobacteria, and Actinobacteria 340

constituted the major parts of biomass on the leaves studied (Figure 3); however 341

a significant proportion of bacteria in the phylloplane did not hybridize to any 342

FISH probe. Coincidentally, these bacteria also exhibited detectable infrared 343

autofluorescence indicative of AAnP (Figures 3) (Atamna-Ismaeel et al., 2012), as 344

which they will be referred to in the following. We found that 2.5% of the 345

autofluorescent bacteria also hybridized to the Alphaproteobacteria-directed 346 Acc

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probe ALF968 and the general probe EUB338. 347

While Alphaproteo-, Betaproteo-, Actinobacteria, and AAnPs were evenly 348

distributed, i.e. they could be found in every FOV of a sample to similar degrees 349

(Figure 3), Bacteroidetes appeared only in some FOVs as cell clusters or 350

individual cells (Figure 2). Gammaproteobacteria were detected rarely and 351

occurred as single cells. 352

353

Bacterial diversity and abundance in the phylloplane of Arabidopsis 354

In general, FISH probes targeting four phylogenetic groups, in combination with 355

the detection of infrared autofluorescence, were sufficient to cover the total 356

bacterial communities. Specific phylogenetic groups covered the phylloplane to 357

different degrees. On average, 1.4% of the phylloplane was covered by 358

Alphaproteobacteria, 2.7% by AAnPs, 0.8% by Betaproteobacteria, 0.4% by 359

Actinobacteria, and 0.2% by Bacteroidetes (Figure 4 A). By adding up the average 360

areas covered by different bacterial groups, i.e. the groups detected by specific 361

FISH and infrared autofluorescence, and comparing the result to the average 362

total phylloplane area covered by bacteria, we found that no major bacterial 363

contributor remained undetected (Figure 4 B). Note, bacteria with an infrared 364

fluorescence that also stained with FISH probes represented a minor fraction 365

and were counted only once. 366

Cell numbers for the analyzed bacterial groups per cm2 leaf surface were 367

estimated by using their median object size (extracted from the segmented 368

images in DAIME and exemplarily validated manually). This approach predicted 369

median areas covered per a cell of 0.5 µm2 for Alphaproteobacteria; 1.4 µm2 for 370

Betaproteobacteria; 0.3 µm2 for Actinobacteria; 1.4 µm2 for Bacteroidetes; and 1.6 371 Acc

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µm2 for AAnPs. The estimated cell densities were between 105 and 2.5 x 106 cm-2 372

per phylogenetic group (Figure 4 C). This conversion from areas to cell numbers 373

shifted the relative abundances of the phylogenetic groups (Figure 4 D and E), as 374

the median cell sizes of the phylogenetic groups differed. Adding up all bacteria 375

we estimate an average bacterial abundance of 5.42 x 106 bacteria per cm2 leaf 376

surface and 1.46 x 108 bacteria per gram fresh weight (leaf fresh weight was 377

estimated from surface area / weight correction factors found in Schmitz 2010). 378

379

Spatial arrangement patterns of phylogenetic groups in the phylloplane of 380

Arabidopsis 381

Based on specific FISH labeling and digital image analysis, it was possible to 382

analyze spatial relationships among the dominating members of the phylloplane 383

community (all images passed the DAIME suitability tests for spatial 384

arrangement analyses). Most analyses indicated co-aggregation of bacteria 385

belonging to different phylogenetic groups at distances below 6 µm (Figure 5). At 386

distances larger than 6 µm, cells were randomly distributed, i.e, pair cross-387

correlation values were not significantly different from 1. Bacteroidetes showed a 388

lower tendency to co-aggregate with members of other groups. Notably, much 389

higher pair correlation values indicated that co-aggregation within phylogenetic 390

groups was much stronger than co-aggregation between groups (Figure 5). 391

Moreover, for bacteria of the same group co-aggregation was in general observed 392

for longer distances (up to 8 µm) than for members of different groups. This 393

indicates that bacteria of the same group preferentially clustered in 394

microcolonies. The tendency of AAnPs to co-aggregate with each other was 395

rather weak as indicated by low pair correlation values at distances beyond the 396 Acc

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median cell size of AAnPs (Figure 5). 397

398

Discussion: 399

FISH approaches have occasionally been used in studies of bacteria colonizing 400

plants (Li et al., 1997; Bisha and Brehm-Stecher, 2009; Bulgarelli et al., 2012) but 401

so far a combinatorial FISH approach that allows uncovering the phylogenetic 402

identity of several phylogenetic groups naturally colonizing the phyllosphere or 403

phylloplane was not performed to our knowledge. 404

It is noteworthy that the bacterial phylogenetic groups identified and their 405

relative abundances at the single-cell level roughly reflect the community 406

structure determined by next generation sequencing approaches (Vorholt, 2012; 407

Bodenhausen et al., 2013), although at lower phylogenetic resolution. Three 408

major phylogenetic groups, namely Alphaproteobacteria, Betaproteobacteria, and 409

Actinobacteria, dominated the phylloplane samples and were evenly distributed. 410

Furthermore, Bacteroidetes were consistently found in every Arabidopsis 411

phylloplane sample observed but appeared only in some FOVs. 412

Gammaproteobacteria were only rarely detected in this study. Since they are 413

nevertheless readily detectable in sizeable proportions in metagenomic studies 414

(Vorholt, 2012) and by 16S rDNA amplicon sequencing (Bodenhausen et al., 415

2013). However, these studies did not distinguish epiphytic from endophytic 416

bacterial populations. It is noteworthy, that the weather had been hot for several 417

weeks prior to sampling, possibly selecting against the fast growing epiphytic 418

Gammaproteobacteria. Moreover, as plant colonizing Gammaproteobacteria are 419

mostly copiotrophs and grow fast when present in "ideal" environments, it is not 420

unlikely that they were clustered in rare hotspots containing high amounts of 421 Acc

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nutrients (Leveau and Lindow, 2001; Monier and Lindow, 2004; Remus-422

Emsermann et al., 2012). 423

The striking abundance of bacteria exhibiting anoxygenic photosystem-related 424

infrared autofluorescence was higher than the predicted relative abundances of 425

10-20% or 6% of the total community in the Arapidopsis phyllosphere based on 426

metagenomes or leaf-wash counts, respectively (Atamna-Ismaeel et al., 2012). 427

Also striking was that none of the applied FISH probes hybridized reliably to 428

these bacteria, and only probes ALF968 and EUB338 occasionally bound to these 429

organisms. We hypothesize that the majority of the AAnPs belonged to the class 430

Alphaproteobacteria (Atamna-Ismaeel et al., 2012). More specifically, they may 431

represent Methylobacterium strains, as it was recently found that all cultivable 432

bacteria exhibiting aerobic anoxygenic photosynthesis in the phyllosphere of 433

white clover were Methylobacteria (Stiefel et al., 2013). Moreover, the cells that 434

we found to exhibit infrared fluorescence were morphologically and 435

phenotypically similar to the pure culture strains shown in Supplemental Figure 436

2 since the observed cells were quite large in comparison to other morphotypes 437

found in the samples and the pure cultures also showed inconsistent 438

hybridization patterns to the aforementioned rRNA-targeted probes. The only 439

other cells that morphologically resembled AAnPs hybridized to the 440

Bacteroidetes-specific probes; however, to our knowledge no Bacteroidetes strain 441

has been described that belongs to the AAnPs. Consistently, none of the 442

Bacteroidetes-related cells exhibited infrared autofluorescence. 443

We observed higher bacterial cell numbers on the abaxial side of the leaves, 444

which is not surprising since the leaf offers more protection on this side. In 445

particular UV light and draught stress (Vorholt, 2012) will be much lower on the 446 Acc

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abaxial side since the leaf blocks much of the UV-fraction of the sunlight and 447

relative humidity can be expected to be higher under plant leaves. 448

We observed aggregation patterns for all probe combinations investigated, with 449

weak but significant cross correlation values at short distances between 450

members of most studied phylogenetic groups (Figure 5). This result indicates a 451

high probability to find bacteria belonging to different phylogenetic groups in 452

close vicinity of each other. However, we interpret this effect as dictated mainly 453

by the environment rather than by bacterial interactions. Bacteria seemed to 454

prevalently proliferate in similar sites like epidermal cell grooves, forcing 455

phylloplane colonizers to grow in close vicinity of each other. Any possible 456

interaction will thus be environmentally amplified due to geographical features 457

of the leaf. We observed significant co-aggregation only with a minimal distance, 458

(see Figure 5) which is in line with observations that multispecies aggregates are 459

not common (Monier and Lindow, 2005b). The much higher probability to find 460

bacteria of the same phylogenetic group to be co-aggregated reflects the growth 461

of microcolonies upon successful colonization. However, these spatial patterns 462

also appeared to be strongly influenced by the leaf environment, since the 463

distance until random distribution occurred was nearly identical for members of 464

the same and different phylogenetic groups (Figure 5). Thus, the habitability of 465

the phylloplane environment appears to shape the bacterial colonization pattern 466

and bacterial life seems to be concentrated at distinct sites in the phyllosphere, 467

where aggregation takes place. Similar effects have been observed previously in 468

bacterial dual species model systems (Monier and Lindow, 2005b). This 469

assumption is in line with a phyllosphere model in which many small sites with 470

intermediate resource availability are prevalent (Remus-Emsermann et al., 471 Acc

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2012). 472

The results presented in this study provide a first assessment of natural 473

community patterns in the phylloplane of the model plant Arabidopsis at a 474

micrometer level and can serve as a basis to investigate the dynamics of 475

phylloplane communities in the future. The methods presented will give us 476

better predictive power for questions on bacterial clustering behavior in situ and 477

may also help to analyze the mechanism of action of plant probiotic bacteria that 478

exhibit a protective effect against pathogenic bacteria (e.g. Innerebner et al., 479

2011; Vogel et al., 2012). It has been suggested that niche occupation is a critical 480

factor contributing to the protective effect known as preemptive colonization 481

(Lindow, 1987; Remus-Emsermann et al., 2013). Therefore, an in-depth spatial 482

description is crucial to understand and interpret such drivers of plant 483

protection. Moreover, the herein developed technique to fix and observe spatial 484

patterns will help to study phylum distributions in the phyllosphere at scales 485

relevant to microbial life with a phylogenetic resolution down to the genus level 486

in the future. 487

488

Acknowledgements: 489

Funding was provided by ETH Zurich and Austrian Science Fund (FWF, grant no. 490

P24101-B22). 491

492

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493 494

References: 495

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673

Table 1: FISH probes applied in this study 674

*competitor probe required 675

676

Figure legends: 677

678

Figure 1: Relative area per FOV covered by bacteria of different samples shown as a 679

normal probability plot. Each point represents the bacterial coverage in one FOV; different 680

symbols correspond to the different samples. No significant differences could be detected 681

between the samples. Horizontal stippled line = average biomass. Vertical stippled line = 682

median and average of the distributions. 683

684

Figure 2: Representative rRNA-targeted FISH micrographs of phylloplane communities 685

sampled by employing the cuticle tape lift technique. A) Green = Actinobacteria; Purple = 686

Betaproteobacteria; Blue = general bacterial probe; AAnPs not shown. Betaproteobacteria 687

are enclosing an imprint of a stomate. B) Purple = Alphaproteobacteria; Green = 688

Betaproteobacteria; Blue = general bacterial probe; AAnPs not shown. Betaproteobacteria 689

are clustering close to a stomate imprint. C) Green = Alphaproteobacteria; Purple = 690

Bacteroidetes; Blue = general bacterial probe; White = AAnPs. Alphaproteobacteria formed 691

Target group Probe name Probe sequence

**competitor probe sequence

Labels Formamide % Reference

Alphaproteobacteria ALF968 GGTAAGGTTCTGCGCGTT Cy3;

Cy5

30 (Neef, 1997)

Betaproteobacteria* Bet42a GCCTTCCCACTTCGTTT **GCCTTCCCACATCGTTT

Cy3;

Cy5

30 (Manz et al., 1992)

Gammaproteobacteria* Gam42a GCCTTCCCACATCGTTT **GCCTTCCCACTTCGTTT

Cy3 30 (Manz et al., 1992)

Bacteroidetes CFB319a TGGTCCGTGTCTCAGTAC Cy3,

Cy5

30 (Manz et al., 1996)

Bacteroidetes CFB719 AGCTGCCTTCGCAATCGG Cy3,

Cy5

30 (Weller et al.,

2000)

Actinobacteria* HGC69A (HGC)

TATAGTTACCACCGCCGT **TATAGTTACCGGCGCCGT

Cy3 30 (Roller et al.,

1994)

Most Bacteria EUB388 GCTGCCTCCCGTAGGAGT Atto488;

5’FAM

20-50 (Amann et al.,

1990)

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many small single layered aggregates. Bacteroidetes formed a small, possibly ball-shaped 692

multilayered microcolony. D) Green = Alphaproteobacteria; Purple = Bacteroidetes; Blue = 693

general bacterial probe; White = AAnPs. Arrows point to stomatal imprints. Scale bars = 10 694

µm 695

696

Figure 3: Relative coverage of leaf samples by different bacterial phylogenetic groups and 697

AAnPs. A = Bacteroidetes; B = Alphaproteobacteria; C = AAnPs; D = Betaproteobacteria; E = 698

Actinobacteria. Every box represents the bacterial coverage in one leaf sample and the 699

recorded FOVs. Box boundaries and middle line represent the 25 percentile, median, and 700

75 percentile; whiskers represent 10 and 90 percentiles, respectively. 701

702

Figure 4: A) Average relative area covered by bacteria B) Average total area covered by 703

bacteria per FOV compared to the average area covered by the specific phylogenetic 704

groups analyzed. Adding up the average occurrence of bacterial groups, we could not 705

detect any differences between the total area covered by bacteria and cumulative 706

abundance of all groups. C) Estimated cell counts of the different groups and AAnPs per 707

cm2 D) Relative area contribution of different groups and E) relative contributions of 708

different groups normalized by cell size. All error bars represent the standard deviation of 709

the mean. 710

711

Figure 5: Analyses of spatial arrangement patterns observed for different phylogenetic 712

groups in the phylloplane and members of the same probe target group. Solid curves show 713

the average pair correlation or cross-correlation values averaged over all images, dotted 714

curves indicate upper and lower 95% confidence limits. The horizontal stippled line 715

indicates values 1, which represents a random spatial distribution. Pair correlation values 716

greater than 1 indicate spatial co-aggregation of the populations analyzed. Values below 1 717

indicate a negative spatial correlation, possibly due to repulsion or niche segregation. The 718

vertical stipple-dashed line represents the intercept of the lower 95% confidence interval 719

with the random distribution line; this point marks the end of a co-aggregation signal and 720 Acc

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the beginning of random spatial distribution. Solid vertical lines in self-correlation plots 721

depict the determined average cell size; pair correlation values >1 at distances larger than 722

the average cell size indicate microcolony formation of bacteria of the same phylogenetic 723

group. 724

725

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31%

8%2%

10%

49%

53%

12% 3%

7%

25%

Actinobacteria

Bacteriodetes

Betaprotebacteria

AlphaproteobacteriaAAnP

total biomass

Are

a co

vere

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]

5

4

3

2

1

0

B

Are

a co

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]

0

1

2

3

4A

4

5

6

7

log

(est

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ount

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cm

2 )

CD E

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spatial distribution analyses of natural phyllosphere-colonizing bacteria on a rabidopsis thaliana...

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