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Applied and Environmental Microbiology, September 2006, p. 6234-6241, Vol. 72, No. 9
0099-2240/06/$08.00+0     doi:10.1128/AEM.00744-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Temporal Changes in Microscale Colonization of the Phylloplane by Aureobasidium pullulans

Molly J. McGrath{dagger} and John H. Andrews*

Plant Pathology Department, University of Wisconsin, Madison, Wisconsin 53706

Received 30 March 2006/ Accepted 19 June 2006


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ABSTRACT
 
Colonization of apple leaves by the yeastlike fungus Aureobasidium pullulans was followed quantitatively and spatially at a microscale level throughout two growing seasons. Ten field leaves were sampled on 11 dates in 2003 and 15 dates in 2004. Using an A. pullulans-specific fluorescence in situ hybridization probe and epifluorescence microscopy, we enumerated total cells, swollen-cells and chlamydospores (SCC), and blastospores/mm2 on leaf features, including the midvein, other (smaller) veins, and the interveinal regions. By 7 July 2003 and 7 June 2004, the total numbers of A. pullulans cells/mm2 were significantly higher (P < 0.05) on the midvein and other veins than in the interveinal regions. This pattern remained consistent thereafter. The primary colonizing morphotype in all regions at all dates was the SCC form, although blastospores always occurred in low numbers. Occupancy was quantified based on the percentage of microscope fields of a particular leaf feature containing ≥1 A. pullulans cell. In general, as seasons progressed, the percent occupancy of features increased and, for most midvein and veinal features, approximated 100% at the end of both growing seasons. Except for early collections, when A. pullulans cell numbers were low, the percent occupancy of interveinal regions was lower than that of the midvein or other veinal regions. A. pullulans was distributed primarily as single cells throughout the seasons in interveinal regions. On the midvein and other veins, colonies of ≥4 cells developed over time, and more cells occurred in colonies than as singletons by August. Our results demonstrate that A. pullulans primarily colonizes veins, where populations appear to increase by growth in situ. This pattern is established early in the growing season and persists.


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INTRODUCTION
 
Microbial ecologists have made quantitative assessments of phylloplane inhabitants for decades, using the whole leaf as a sampling unit. These studies have generated a wealth of information on seasonal changes in phylloplane communities (30), positional variation in phylloplane populations within a canopy (3), the influence that population size has on plant disease (15, 18), the impact of population dynamics on phylloplane communities (13), and the abilities of potential biocontrol organisms to survive under stressful environmental conditions (19, 33). Such data have contributed greatly to the understanding of phylloplane microbial ecology. Unfortunately, what is not addressed in most of these studies is colonization under field conditions at the microscale level, which is the scale that corresponds to the size of a microbe (24). If colonization of the leaf surface were uniform, the leaf in its entirety would be representative for studies of phylloplane population size. However, microbes often have been found in characteristic patterns clustered along veins, in grooves between epidermal cells, or at the bases of trichomes (4, 16, 20). Consequently, important information about microbial colonization processes is likely lost wherever the whole leaf is a sampling unit.

Microscale colonization was examined recently by tools such as a specific fluorescence in situ hybridization (FISH) probe and epifluorescence microscopy, and Aureobasidium pullulans was found to form cell aggregates on the midvein, other veins, and wounds on the apple phylloplane (4). However, sampling was periodic, and it is unclear when this pattern arises and if it persists throughout the growing season. Possibly it does not persist because the physical surface of the leaf is highly dynamic. The cuticle can erode progressively, which can change the topography, the wettability of the surface, and exudation of nutrients (22, 27, 28). The colonizing microbial community is also dynamic, and some inhabitants may impact other colonists by producing inhibitory compounds (21) or by facilitating colonization (23). Consequently, phylloplane colonizers may constantly be forced to compete for sites and resources on a changing leaf surface (5, 21).

Here, we extend the use of our model system to quantify A. pullulans and to document the spatial pattern of A. pullulans populations at frequent intervals over the course of an entire growing season. A. pullulans, a deuteromycete black yeast (7), is a ubiquitous phylloplane inhabitant with potential as a biocontrol agent of foliar pathogens, such as Venturia inaequalis (1). It is polymorphic, with morphotypes that include blastospores, swollen cells and chlamydospores (SCC), pseudohyphae, and hyphae (26). The fungus produces extracellular polysaccharide (EPS), which promotes adhesion and may prevent desiccation and/or provide protection from environmental stresses (2, 32).

We were interested in three population parameters: abundance (the number of individual A. pullulans cells in an area), occupancy (the occurrence of A. pullulans in a given area, typically a microscope field of a leaf feature), and colonization (establishment of the A. pullulans population on a leaf feature) (9-11). These are fundamental ecological parameters that tend to be related (10). For example, as overall abundance declines, the number of occupied sites also declines; conversely, as abundance increases, the number of occupied sites also generally increases (9). This relationship has been described as one of the most important in ecology (9). However, the nature of the relationship remains unclear, including the roles of spatial and temporal scales (9). Since resource availability can vary temporally and spatially (8), the presence of colonies can reveal whether sites can support growth. This is the first time these parameters have been studied for a microbial species over a significant period of time (an entire growing season) at the microscale level on field leaves. Such information should enhance our understanding of the colonization process in general (10), as well as its relevance to phyllosphere microbial ecology in particular.

The objectives of this study were specifically to determine (i) when microscale colonization patterns develop on field leaves; (ii) if these patterns, once formed, change over the course of the growing season; (iii) the prevalent A. pullulans morphotypes over time and on individual features; (iv) if occupancy of individual features changes over time; (v) the distribution of A. pullulans cells in colonies on features; and (vi) if Geographical Information System (GIS) maps can be used as a rapid method to visually assess critical changes in the distribution of a microbe at a microscale level.


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MATERIALS AND METHODS
 
Leaf collections.
Experiments were conducted in 2003 and 2004 with apple leaves from terminal shoots (Malus domestica cv. Liberty scions grafted to M26 rootstock) taken from saplings planted in 1998. The trees were approximately 2.5 to 3 m tall, untreated with fungicide, and located at the West Madison Agricultural Research Station in Madison, WI, in four blocks among three other apple cultivars, also in blocks. Ten leaves at each sampling date were chosen by assigning numbers to 22 trees and randomly generating numbers with replacement (corresponding to apple trees) to select the trees from which sample leaves were taken on the following dates in 2003: 3 May; 16 and 30 June; 7, 14, 21, and 29 July; 4 and 15 August; and 16 and 22 September. In 2004, the sampling dates were 3, 10, and 18 May; 7, 14, and 28 June; 7, 12, 19, and 26 July; 6 and 16 August; 7 and 30 September; and 12 October. Average-size leaves on the west side of the designated trees about 1.5 to 2 m from the orchard floor were removed at the base of the petiole. Individual leaves were placed in sterile Whirl-Pak (Nasco, Fort Atkinson, WI) bags and transported in an insulated cooler with ice to the laboratory for processing.

Sample processing.
Three segments approximately 5 mm wide were removed from the apical third, middle, and basal third of each leaf. If the leaf was too small to remove three 5-mm segments, the entire leaf was processed. Leaf tissue was fixed in 3:1 methanol-glacial acetic acid and stored in 70% ethanol at –20°C until it was processed. Segments were then removed from storage, washed, and prepared for FISH as described previously (17, 29). The A. pullulans-specific FISH probe detects all forms of A. pullulans on leaves (17). Every microscope field along two nonadjacent transects per segment (one about 1.5 mm from the tip and the other about 1.5 mm from the base of the segment) was viewed with epifluorescence microscopy with a 40x long-working-distance objective lens (area of field, 0.196 mm2). Where whole (small) leaves were processed, six evenly spaced, nonadjacent transects were taken.

Data recording.
A. pullulans cell counts were recorded by morphotype (blastospore, SCC, or hyphal) (26) and degree of aggregation (single cells or colony size) with respect to the leaf region (midvein, other smaller veins, and interveinal). Blastospores and SCC were distinguished based on characteristic morphology (cell size and pigmentation) (2, 25, 26). A field containing the midvein was designated a midvein field, and all A. pullulans cells in the field were considered occupants of a midvein region. Fields containing smaller veins (secondary, tertiary, or finer orders) were designated "other veinal fields," and all cells within that field were considered to be occupying "other (smaller) veinal regions." Finally, fields with neither smaller veins nor the midvein were counted as interveinal fields, and A. pullulans cells in these fields were designated occupants of the interveinal regions. Unoccupied fields were similarly recorded, so the number of cells per mm2 of leaf area containing a region of interest could be calculated. Cells in direct contact were considered to be a colony. Multiple morphotypes and colonies and/or single cells were occasionally present in the same field.

Statistical analysis.
A. pullulans population sizes were log10 (x + 1) transformed to achieve normality prior to statistical analysis. To determine if differences in numbers of cells per mm2 on midveins and other veins and in interveinal regions existed, analysis of variance tests were performed with the PROC MIXED procedure in SAS version 8.2 (SAS Institute, Cary, N.C.). This allowed all regions to be compared (midvein to other veins, midvein to interveins, and other veins to interveins) at all dates. The PROC MIXED procedure was also used to determine if SCC and blastospore densities differed in the individual regions. The percentages of occupation of individual regions were also compared by using the PROC MIXED procedure. A microscope field was considered occupied if it contained ≥1 A. pullulans cell. Percent occupancy by feature was calculated as the number of occupied fields containing the feature of interest divided by the total number of fields containing that feature. To determine if the percent occupation of the total leaf differed over time, Student's t tests (assuming unequal variances) were conducted by using Microsoft Excel 2003 v. 11 (Redmond, WA). The percentages of individual A. pullulans cells distributed as single cells and colonies of 2 or 3, 4 to 9, or ≥10 cells were also compared by using Student's t test (assuming unequal variances) with Microsoft Excel 2003 v. 11. Colony (or single-cell) sizes were compared among the individual features on the same sampling dates.

GIS plots.
To facilitate the interpretation of colonization patterns, ArcView GIS version 3.2 (ESRI, Redlands, CA) was used to represent spatial data visually. Macro programs were generated by using Microsoft Excel, which stacked transects for GIS plots on a prime meridian with the midvein as a vertical reference point (4). This allowed the cell number, cell type, and leaf region for each microscope field to be placed on an artificial grid of longitude and latitude. Leaves (one per date) from 3 May, 7 June, and 30 September 2004, representative of A. pullulans cells, colony size, and distribution on the respective dates, were chosen for the GIS plots to depict the major trend in colonization. Spatial data from three segments and two transects per leaf were used for each GIS plot.


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RESULTS
 
Pattern development on field leaves.
Low numbers of A. pullulans cells were detected on midveins and other veins and in interveinal regions of field leaves as early as 3 May 2003 and 3 May 2004 (Fig. 1A and B). There was no established pattern of differences with respect to colonization by feature until the 7 July 2003 sampling date. On that date and all sampling dates thereafter, the trend did not change: all regions were significantly different (P ≤ 0.05) from each other, with A. pullulans densities on midveins higher than on other veins and in interveinal areas and densities over other veins higher than over the interveinal regions.


Figure 1
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  		FIG. 1. Mean log10 (x + 1) A. pullulans (Ap) cells/mm2 (± standard deviation) on midveins and other veins and in interveinal regions over the 2003 (A) and 2004 (B) growing seasons. Sampling dates are identified by the month-year convention. The numbers of cells/mm2 were determined by using the total number of fields (each field was 0.196 mm2) per leaf transect containing the feature of interest (midveins, other veins, or intervein). Ten leaves were examined on each sampling date. Three segments/leaf and two transects/segment were evaluated. The arrows indicate the first date on which all three regions were significantly different (P < 0.05) from each other.

Early in 2004, there was no established colonization pattern until 7 June (Fig. 1B). At that date, the pattern stabilized in the same relative order as for 2003 and remained in that order thereafter.

Morphotype of A. pullulans.
The results in 2003 and 2004 were similar; thus, only data for 2004 are presented (Fig. 2A to C). Blastospores were always found on at least one site on all sampling dates, but the numbers of blastspore cells were extremely low at all times. The predominant morphotype was SCC. The numbers of SCC per mm2 on midveins and other smaller veins and in interveinal regions were significantly higher (P ≤ 0.05) than the numbers of blastospores on each respective leaf feature at all sampling dates. By 7 June and on all sampling dates thereafter, SCC densities were significantly higher over midveins than either over other veins or in interveinal regions, and densities over other veins were significantly higher than those over interveinal features.


Figure 2
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  		FIG. 2. Mean log10 (x + 1) A. pullulans (Ap) SCC and blastospores/mm2 (± standard deviation) on midveins (A) and other veins (B) and in interveinal regions (C) over the 2004 growing season. Sampling dates are identified by the month-year convention. The numbers of cells per mm2 were determined by using the total number of microscope fields (each field was 0.196 mm2) per leaf transect containing the feature of interest (midveins, other veins, or interveinal sites). Fields included one or both morphotypes (SCC and blastospores). Ten leaves were examined on each sampling date. Three segments/leaf and two transects/segment were evaluated.

Occupation of available leaf features by A. pullulans.
The results in the 2003 and 2004 growing seasons were similar, so only the 2004 data are summarized here. The percentages of occupied microscope fields, overall, on the first four sampling dates (3 May to 7 June) were significantly lower (P ≤ 0.05) than the percentages of occupation on all other sampling dates (Table 1). Commencing on 14 June, at least 50% of the overall fields were occupied, with the highest occupancy occurring on 6 August, when about 80% of all fields contained ≥1 A. pullulans cell.


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  		TABLE 1. Occupancy of individual leaf features (midvein, other vein, or interveinal) and the overall leaf by Aureobasidium pullulans cells

With respect to leaf features, the percentage of midvein fields occupied by one or more A. pullulans cells increased steadily with time (Table 1). Beginning relatively early (7 June), all midvein fields were occupied. On 3 May and thereafter, the percentages of occupied midvein fields were significantly higher (P ≤ 0.05) than the percentages of occupied smaller vein or interveinal fields (with the exception of 6 August, 7 September, and 12 October, when the percentages of occupied midveins and other veins were not significantly different). On all sampling dates after 18 May, the percentage of occupied smaller vein fields was significantly higher (P ≤ 0.05) than the percentage of interveinal fields occupied. The largest percentages of occupied smaller-vein fields were about 98% on 6 August and 12 October. On the first two sampling dates only (3 May and 10 May), the percentages of interveinal fields occupied were significantly higher (P ≤ 0.05) than the percentages of occupied smaller-vein fields (there was not a significant difference between other veins and interveinal fields on May 18). The percent occupation of interveinal fields fluctuated substantially throughout the growing season but overall averaged less than 50%.

Colony formation.
The 2003 and 2004 results were similar, so only the 2004 results are presented. Early in 2004 (3 May and 10 May), A. pullulans was distributed in all regions largely as single cells (Fig. 3A to C). A. pullulans continued to be distributed primarily (≥70% of A. pullulans cells) as singletons in interveinal regions (Fig. 3C). Fewer cells were in colonies of 2 or 3 cells (no more than 22% at any date), and very few A. pullulans cells were in colonies of 4 to 9 or ≥10 cells (less than 5% on all sampling dates). On midveins and other smaller veins, the percentages of A. pullulans cells in colonies increased over time (Fig. 3A and B). Few cells (P ≤ 0.05) were distributed as individuals on midveins and other smaller veins on 14 June and thereafter compared to those on interveinal features. At the end of the season, a higher percentage of cells were distributed in colonies of ≥10 cells (over 20% of cells on some dates on midveins and over 10% on some dates on other veins). By 6 August and thereafter, significantly more cells (P ≤ 0.05) were distributed in the largest colonies (colonies of 4 to 9 and ≥10) on midveins and smaller veins than in interveinal regions (with the exception of 12 October, when there were no significant differences between the numbers of colonies of ≥10 cells on other veins and in interveinal regions). The highest mean numbers of cells in colonies of four to nine cells/mm2 on midveins and other veins and in interveinal regions were 104.6 ± 19.7 (on 16 August), 41.4 ± 14.8 (6 August), and 2.7 ± 3.2 (6 August), respectively (data not shown). The highest mean numbers of colonies of ≥10 cells/mm2 on midveins and other veins and in interveinal regions were 62.8 ± 41.6 (30 September), 12.6 ± 11.1 (6 August), and 0.026 ± 0.081 (7 September), respectively (data not shown). While colonies were numerous on other veins, and especially midvein sites, at least some portions of veins in all fields remained uncolonized.


Figure 3
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  		FIG. 3. Percentages of A. pullulans (Ap) cells distributed as single cells (horizontal stripes) and in colonies of 2 or 3 cells (dark gray), 4 to 9 cells (checkered), or ≥10 cells (black) on midveins (A) and other veins (B) and in interveinal regions (C) during the 2004 growing season. Sampling dates are identified by the month-year convention. Colony size was defined as the number of A. pullulans cells in direct contact. Each field was designated as midvein, other vein, or interveinal, but multiple colonies and/or single cells could be present in each field.

GIS plots.
Plots were generated from single leaves sampled at three key dates in 2004. The first plot (Fig. 4), from a leaf sampled on 3 May, displays 149 fields from six transects (6 midvein, 44 other vein, and 99 interveinal fields), showing that there were low numbers of A. pullulans cells of both morphotypes and that most cells were distributed on interveinal sites. The second plot (Fig. 5) (from 7 June) depicts the time at which the prevalent veinal colonization pattern became established. This plot for 359 fields (6 midvein, 101 other vein, and 252 interveinal fields) on six transects shows increased numbers of SCC, while numbers of blastospore cells remained low. Though there were about the same quantities of A. pullulans cells on interveinal sites and other veins, the interveinal regions made up about 70% of the total fields. Consequently, the numbers of cells/mm2 on other veins were significantly higher (P ≤ 0.05) than in interveinal regions, and occupancy was higher (Table 1). The last plot (Fig. 6) (from 30 September) is representative of the number of A. pullulans cells and distribution late in the growing season, when the colonization pattern (visible in the 7 June plot) was even more apparent. The plot displays 469 total fields (8 midvein, 126 other vein, and 335 interveinal fields) on six transects. In both distribution (occupancy of microscope fields) and density, cells (nearly all SCC) were highly localized to veins.


Figure 4
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  		FIG. 4. GIS plot showing the spatial distribution of Aureobasidium pullulans on the adaxial surface of an apple leaf sampled on 3 May 2004, when A. pullulans SCC and blastospore counts were low. The leaf is representative of the 10 leaves sampled and examined by FISH on that date. Three leaf segments (separated vertically by blank space) and two transects per segment are displayed. All microscope fields (one field was 0.196 mm2) from each transect are depicted. The top line of the transect maps the blastospore counts, the middle line displays the SCC counts, and the bottom line shows the microsite features present in the fields.


Figure 5
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  		FIG. 5. GIS plot showing the spatial distribution of Aureobasidium pullulans on the adaxial surface of an apple leaf sampled on 7 June 2004, when a veinal pattern in A. pullulans density began to develop (see the legend to Fig. 4 for GIS plot specifications).


Figure 6
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  		FIG. 6. GIS plot showing the spatial distribution of Aureobasidium pullulans on the adaxial surface of an apple leaf sampled on 30 September 2004, when A. pullulans cells were primarily SCC and highly localized to veinal sites (see the legend to Fig. 4 for GIS plot specifications).


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DISCUSSION
 
To our knowledge, this is the first comprehensive microscale assessment of seasonal changes in microbial distribution and colonization of the phylloplane, though other studies have examined spatial issues under various permutations of controlled conditions, for much shorter periods of time, or with less spatial precision (6, 16). Andrews et al. (4) sampled apple leaves on eight dates in 1998 and 1999 and determined that A. pullulans was distributed primarily on veins (and microwounds). Here, sampling throughout entire growing seasons allowed us to determine when the colonization pattern, like that observed in 1998 and 1999, develops and if temporal changes in colonization occur. In turn, this information should illuminate the explanation for the pattern and its biological significance.

One important result from this study relates to the onset of veinal colonization. Early (May and June) in both seasons, the occupancy of the midvein, other veins, and interveinal regions appeared to be random, likely the result of random immigration events. This was reflected in low numbers of cells of A. pullulans on all features, where on some transects there were no A. pullulans cells on veinal sites. Our evidence also suggests that A. pullulans arrives as single cells (at least early in the season), because nearly all cells at this time were singletons. Immigration followed by successful establishment constitutes colonization (11). High immigrant numbers would increase the chances of an individual immigrant arriving at a site favorable for growth. Significant differences in spatial pattern then become apparent. The source and size of the A. pullulans immigrant pool throughout the growing season need to be investigated, along with whether young leaves can support the formation of colonies.

As the season progressed, large colonies formed mainly on veinal sites. This shows that the leaf surface as it develops can support at least local growth with endogenous or possibly exogenous nutrients. Intensive, season-long sampling showed that veins evidently also support more growth late in the growing season, as reflected by the presence of larger and more numerous A. pullulans colonies over time. Colony sizes were consistently larger on veins, but there were occasionally small colonies in interveinal regions, so more or less growth was possible on all features. Dense colonization of veins has been attributed to preferential accumulation of cells in veinal channels due to rain events (25, 31) or to growth promoted by higher nutrient concentrations (16, 25).

While large colonies were present on veins, and almost all (95%) veinal fields were declared occupied (i.e., they contained at least one A. pullulans cell) at the end of the season, we observed uninhabited areas on all veins, even on the densely colonized midvein. Possibly only portions of these veins can support A. pullulans growth, either due to the differential presence or absence of growth-promoting or -inhibiting factors or because of prior occupation by other microorganisms. Alternatively, veinal colonies may expand slowly because organisms on the phylloplane are subjected to periodic harsh environmental conditions. Some evidence for this possibility is provided by the finding that SCC were the dominant morphotype at all times on all sites in the growing season. This is the morphotype that produces EPS, which is thought to promote adhesion, prevent desiccation, and/or provide protection from environmental stresses, such as UV radiation (2, 32). The outer wall of chlamydospores contains melanin (26), and pigments have also been shown to provide protection from UV radiation (12).

SCC may be the predominant morphotype, but blastospores were present in low numbers on all sampling dates. It is speculated that they are the dispersal form, because they lack EPS and are the budded morphotype (2, 26). However, their paucity indicates that if they are the immigrating morphotype, they likely convert quickly on the phylloplane to SCC (the probable survival form), die, or are eroded from the leaf by wind or rain. When conditions are favorable for growth, SCC apparently bud blastospores, forming colonies of the sizes observed late in the growing season. Further studies are needed to determine the biotic and abiotic factors that trigger growth episodes.

The GIS plots provided a rapid, visual method to express and interpret spatial colonization data. These maps could be extended to express similar data for other epiphytes or pathogens, thereby depicting the topographical data for competitors. Such displays could be useful in screening for prospective antagonists (6).

Kinkel et al. (14) proposed a general framework for microbial colonization of leaves that is relevant to some of our findings. According to their model, aggregation of resources would reduce the total number and proportion of colonizable sites when immigration is random. We found that colonization was highly localized to veinal regions, likely, according to the model, because resources are more available in those areas. Another prediction is that when resources are aggregated, immigrants to these sites will have a greater local reproductive potential than if resources are distributed uniformly. Large A. pullulans colonies in veinal regions, and their absence on interveinal features, fit this prediction. If this is the case, there is also a greater mean carrying capacity per colonizable feature (in the absence of saturating immigration), and it would take longer or more frequent periods of growth-promoting conditions for a microbial population to reach carrying capacity than if resources were uniformly distributed. Our results showed that veinal regions were not fully colonized, possibly because there are only fleeting periods during which conditions allow population expansion. If resources are aggregated, in the absence of immigrants some of the favorable microsites are unoccupied, and the leaf remains below its carrying capacity. This seemed to be evident early in the season, when A. pullulans cell counts were low. As the season progressed, a greater percentage of the leaf area became occupied. Growth occurs during occasional favorable periods by the budding of blastospores from SCC. Concurrently, new immigrants arrive and allow further colonization of the veins. The interveinal area appears to be a relatively inhospitable and largely unoccupied terrain, where little if any growth occurs.


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ACKNOWLEDGMENTS
 
Support by NSF grant DEB-0075358 and USDA Hatch grants 142-P446 and 142-4873 is acknowledged with appreciation.

We thank Russell Spear for helping in many ways and Jack Whitney and Keyton Kuykendall for laboratory and field assistance.


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FOOTNOTES
 
* Corresponding author. Mailing address: Plant Pathology Department, 1630 Linden Dr., University of Wisconsin, Madison, WI 53706. Phone: (608) 262-9642. Fax: (608) 263-2626. E-mail: jha{at}plantpath.wisc.edu. Back

{dagger} Present address: Plant Pathology Department, Michigan State University, East Lansing, MI 48824. Back


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REFERENCES
 
    1
  1. Andrews, J. H., F. M. Berbee, and E. V. Nordheim. 1983. Microbial antagonism to the imperfect stage of the apple scab pathogen, Venturia inaequalis. Phytopathology 73:228-234.
    2. 2
  2. Andrews, J. H., R. F. Harris, R. N. Spear, G. W. Lau, and E. V. Nordheim. 1994. Morphogenesis and adhesion of Aureobasidium pullulans. Can. J. Microbiol. 40:6-17.
    3. 3
  3. Andrews, J. H., C. M. Kenerley, and E. V. Nordheim. 1980. Positional variation in phylloplane microbial populations within an apple tree canopy. Microb. Ecol. 6:71-84.[CrossRef]
    4. 4
  4. Andrews, J. H., R. N. Spear, and E. V. Nordheim. 2002. Population biology of Aureobasidium pullulans on apple leaf surfaces. Can. J. Microbiol. 48:500-513.[CrossRef][Medline]
    5. 5
  5. Brodie, I. D. S., and J. P. Blakeman. 1975. Competition for carbon compounds by a leaf surface bacterium and conidia of Botrytis cinerea. Physiol. Plant Pathol. 6:125-135.
    6. 6
  6. Collins, D. P., B. J. Jacobsen, and B. Maxwell. 2003. Spatial and temporal population dynamics of a phyllosphere colonizing Bacillus subtilis biological control agent of sugar beet Cercospora leaf spot. Biol. Control 26:224-232.[CrossRef]
    7. 7
  7. De Hoog, G. S., and M. R. McGinnis. 1987. Ascomycetous black yeasts, p. 187-199. In G. S. de Hoog, M. T. Smith, and A. C.M Weijman (ed.), The expanding realm of yeast-like fungi. Elsevier Science Publishers, New York, N.Y.
    8. 8
  8. Dyer, A. R., and K. J. Rice. 1999. Effects of competition on resource availability and growth of a California bunchgrass. Ecology 80:2697-2710.[CrossRef]
    9. 9
  9. Gaston, K. J., T. M. Blackburn, J. J. D. Greenwood, R. D. Gregory, R. M. Quinn, and J. H. Lawton. 2000. Abundance-occupancy relationships. J. Appl. Ecol. 37(Suppl. 1):39-59.
    10. 10
  10. He, F., and K. J. Gaston. 2000. Estimating species abundance from occurrence. Am. Nat. 156:553-559.[CrossRef]
    11. 11
  11. Ims, R. A., and N. G. Yoccoz. 1997. Studying transfer processes in metapopulations, p. 247-265. In I. Hanski and M. E. Gilpin (ed.), Metapopulation biology. Academic Press, San Diego, Calif.
    12. 12
  12. Jacobs, J. L., T. L. Carroll, and G. W. Sundin. 2005. The role of pigmentation, ultraviolet radiation tolerance, and leaf colonization strategies in the epiphytic survival of phyllosphere bacteria. Microb. Ecol. 49:104-113.[CrossRef][Medline]
    13. 13
  13. Kinkel, L. L., J. H. Andrews, and E. V. Nordheim. 1989. Microbial introductions to apple leaves: influences of altered immigration on fungal community dynamics. Microb. Ecol. 18:161-173.[CrossRef]
    14. 14
  14. Kinkel, L. L., M. R. Newton, and K. J. Leonard. 2002. Resource aggregation in the phyllosphere: implications for microbial dynamics across spatial scales, p. 317-340. In S. E. Lindow, E. I. Hecht-Poinar, and V. J. Elliot (ed.), Phyllosphere microbiology. APS Press, St. Paul, Minn.
    15. 15
  15. Kinkel, L. L., M. Wilson, and S. E. Lindow. 2000. Plant species and plant incubation conditions influence variability in epiphytic bacterial population size. Microb. Ecol. 39:1-11.[CrossRef][Medline]
    16. 16
  16. Leveau, J. H. J., and S. E. Lindow. 2000. Appetite of an epiphyte: quantitative monitoring of bacterial sugar consumption in the phyllosphere. Proc. Natl. Acad. Sci. USA 98:3446-3453.
    17. 17
  17. Li, S., D. Cullen, M. Hjort, R. N. Spear, and J. H. Andrews. 1996. Development of an oligonucleotide probe for Aureobasidium pullulans based on the small-subunit rRNA gene. Appl. Environ. Microbiol. 62:1514-1518.[Abstract]
    18. 18
  18. Lindow, S. E. 1987. Severity of pear fruit russetting associated with epiphytic indolacetic acid-producing bacteria. Phytopathology 77:1724.
    19. 19
  19. Lindow, S. E. 1991. Determinants of epiphytic fitness in bacteria, p. 295-314. In J. H. Andrews and S. S. Hirano (ed.), Microbial ecology of leaves. Springer-Verlag, New York, N.Y.
    20. 20
  20. Mansvelt, E. L., and M. J. Hattingh. 1989. Scanning electron microscopy of colonization of pear leaves by Pseudomonas syringae pv. syringae. Can. J. Bot. 65:2517-2522.
    21. 21
  21. McCormack, P. J., H. G. Wildman, and P. Jeffries. 1994. Production of antibacterial compounds by phylloplane inhabiting yeasts and yeastlike fungi. Appl. Environ. Microbiol. 60:927-931.[Abstract/Free Full Text]
    22. 22
  22. Mechaber, W. L., D. B. Marshall, R. A. Mechaber, R. T. Jobe, and F. S. Chew. 1996. Mapping leaf surface landscapes. Proc. Natl. Acad. Sci. USA 93:4600-4603.[Abstract/Free Full Text]
    23. 23
  23. Monier, J. M., and S. E. Lindow. 2005. Aggregates of resident bacteria facilitate survival of immigrant bacteria on leaf surfaces. Microb. Ecol. 49:343-352.[CrossRef][Medline]
    24. 24
  24. Morris, C. E., J. Monier, and M. Jacques. 1997. Methods for observing microbial biofilms directly on leaf surfaces and recovering them for isolation of culturable microorganisms. Appl. Environ. Microbiol. 63:1570-1576.[Abstract]
    25. 25
  25. Pugh, G. J. F., and N. G. Buckley. 1971. The leaf surface as a substrate for colonization by fungi, p. 431-445. In T. F. Preece and C. H. Dickinson (ed.), Ecology of leaf surface micro-organisms. Academic Press, New York, N.Y.
    26. 26
  26. Ramos, S., and A. I. Garcia. 1975. A vegetative cycle of Pullularia pullulans. Trans. Br. Mycol. Soc. 64:129-135.
    27. 27
  27. Schönherr, J., and P. Baur. 1996. Cuticle permeability studies, p. 1-23. In C. E. Morris, P. C. Nicot, and C. Nguyen-The (ed.), Aerial plant surface microbiology. Plenum Press, New York, N.Y.
    28. 28
  28. Schreiber, L., M. Skrabs, K. D. Hartmann, P. Diamantopoulos, E. Simanova, and J. Santrucek. 2001. Effect of humidity on cuticular water permeability of isolated cuticular membranes and leaf disks. Planta 214:274-282.[Medline]
    29. 29
  29. Spear, R. N., S. Li, E. V. Nordheim, and J. H. Andrews. 1999. Quantitative imaging and statistical analysis of fluorescence in situ hybridization (FISH) of Aureobasidium pullulans. J. Microbiol. Methods 35:101-110.[CrossRef][Medline]
    30. 30
  30. Thompson, I. P., M. J. Bailey, J. S. Fenlon, T. R. Fermor, A. K. Lilley, J. M. Lynch, P. J. McCormack, M. P. McQuilken, K. J. Purdy, P. B. Rainey, and J. M. Whipps. 1993. Quantitative and and qualitative seasonal changes in the microbial community from the phyllosphere of sugar beet (Beta vulgaris). Plant Soil 150:177-191.[CrossRef]
    31. 31
  31. Wilson, D., and G. C. Carroll. 1994. Infection studies of Discula quercina, an endophyte of Quercus garryana. Mycologia 86:635-647.
    32. 32
  32. Wilson, H. A., V. G Lilly, and J. G. Leach. 1965. Bacterial polysaccharides. IV. Longevity of Xanthomonas phaseoli and Serratia marcescens in bacterial exudates. Phytopathology 55:1135-1138.[Medline]
    33. 33
  33. Wilson, M., S. S. Hirano, and S. E. Lindow. 1999. Location and survival of leaf-associated bacteria in relation to pathogenicity and potential for growth within the leaf. Appl. Environ. Microbiol. 65:1435-1443.[Abstract/Free Full Text]

Applied and Environmental Microbiology, September 2006, p. 6234-6241, Vol. 72, No. 9
0099-2240/06/$08.00+0     doi:10.1128/AEM.00744-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • McGrath, M. J., Andrews, J. H. (2007). Role of Microbial Immigration in the Colonization of Apple Leaves by Aureobasidium pullulans. Appl. Environ. Microbiol. 73: 1277-1286 [Abstract] [Full Text]  

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