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© 2013 Plant Management Network.
Accepted for publication 3 April 2013. Published 23 August 2013.

Sporulation Capacity and Longevity of Puccinia horiana Teliospores in Infected Chrysanthemum Leaves

Morris R. Bonde, Cristi L. Palmer, Douglas G. Luster, Susan E. Nester, Jason M. Revell, and Dana K. Berner, United States Department of Agriculture, Agricultural Research Service, Foreign Disease-Weed Science Research Unit, Fort Detrick, Frederick, MD 21702-5023; and the IR-4 Project, Rutgers University, Princeton, NJ 08520

Corresponding author: Morris R. Bonde.

Bonde, M. R., Palmer, C. L., Luster, D. G., Nester, S. E., Revell, J. M., and Berner, D. K. 2013. Sporulation capacity and longevity of Puccinia horiana teliospores in infected chrysanthemum leaves. Online. Plant Health Progress doi:10.1094/PHP-2013-0823-01-RS.


Puccinia horiana Henn., a quarantine-significant fungal pathogen and causal agent of chrysanthemum white rust (CWR), was first discovered in the United States in 1977 and later believed to have been eradicated. Recently, however, the disease has sporadically reappeared in the northeastern US. Possible explanations for the reappearance include survival of the pathogen in the local environment, and reintroduction from other locations. To determine the possibility that the pathogen might be overwintering in the field, we undertook the study described here. Results from the study showed that P. horiana teliospores, imbedded in infected leaves, were capable of sporulating 2 weeks after inoculation, and this capacity continued until the leaf became necrotic and desiccated. This is the first report of the extreme susceptibility of P. horiana teliospores to leaf necrosis and desiccation and suggests that field infections following winter are unlikely to originate from teliospores. Teliospore germination on excised leaves was shown to be inhibited by light.


Puccinia horiana Henn., causal agent of chrysanthemum white rust (CWR), is a quarantine-significant fungal pathogen in the United States. It is indigenous to Asia (11), and was first discovered in Japan in 1895 and described by Hennings (10) in 1901. The disease has been reported in several European countries (1), the United Kingdom (1), New Zealand (8), South Africa (8), Australia (7), South America (6), and states within the US including Pennsylvania (6) and New Jersey (14).

By the late 1970s, CWR appeared to have been eradicated in the US, but reappeared in Washington and Oregon in 1990 (9). In each instance, the disease was detected in hobbyist plantings and quickly eradicated (2). In 1991 and 1992, CWR again was discovered in the US, this time in several counties in California where it caused considerable concern to the nursery industry (3,4,5). As a result, the USDA-ARS Foreign Disease-Weed Science Research Unit, Fort Detrick, MD, in cooperation with the USDA-APHIS, conducted a study to develop a treatment protocol to control the disease (2). From that research, the systemic fungicide myclobutanil was shown to have high curative properties and was adopted by APHIS as a mandatory method to control the disease when discovered (2).

For the next several years, the disease was effectively managed. Recently, however, CWR has reemerged on several occasions, including in Pennsylvania from 2004 to 2012 (12,13). Kim et al. (12) and O’Keefe and Davis (13) provided evidence suggesting the pathogen might be surviving through the winter in the field on infected chrysanthemum plants.

P. horiana is an autoecious microcyclic rust fungus, thus completing its life cycle on a single host. Although the pathogen can infect more than 10 chrysanthemum species, primarily it is known as a pathogen of Chrysanthemum × morifolium. P. horiana produces teliospores that are embedded in infected leaves (8). They germinate to form basidiospores that are the primary infectious propagule (8). Urediniospores are not known to be produced. The amount of sporulation (basidiospores) arising from telia can be extremely high on infected plants, causing the leaf to turn white, hence the name of the disease. Basidiospores are susceptible to desiccation and survive less than 5 min at 80% RH, and less than 1 h at 90% RH.

Teliospores of P. horiana have been shown to survive up to 8 weeks in detached leaves at 50% RH, and for shorter durations at higher relative humidities (8). When buried in compost, they survived for a maximum of 2 weeks, and in moist soil 1 week (8). However, the single report describing teliospore survival relied on methodology not conducive for conclusive determination of teliospore viability at very low levels, which is needed for regulatory purposes (8).

The role of teliospores as a means of long-term survival of P. horiana is not clear, as is the duration teliospores can remain viable and able to sporulate in infected plants, releasing basidiospore inoculum into the environment. The objectives of this study were to: (i) develop a better method to reliably detect and measure viability of P. horiana teliospores in infected chrysanthemum leaves; and (ii) use that method to determine the relationship of pustule and teliospore age to basidiospore production. Information gained would benefit determination of the role of teliospores in survival of the pathogen, and development of CWR epidemics.

Relationship of Leaf Position and Lesion Density to Sporulation

Since P. horiana, being an obligate pathogen, inoculation of test plants requires transfer of inoculum from infected plants to recipient plants. Two chrysanthemum test plants were inoculated by placing them alongside donor plants in a 1.15-m long × 0.84-m wide × 0.95-m high mist chamber that provided a fine mist and chamber RH of 100%. The use of a mist chamber with infected donor plants interspersed with recipient plants provided the appropriate amount of moisture and air turbulence for sporulation on infected leaves, distribution of inoculum, and initiation of infection on recipient chrysanthemum plants. Following approximately 24 h in the mist chamber, recipient plants were placed on a greenhouse bench at 22 to 26°C for disease development. After 7 to 10 days, inoculated plants were examined for symptoms of infection.

In order to measure sporulation capability, we developed a method to quantify basidiospores released from individual infected leaves over extended periods of time. Leaves with pustules from two infected plants were collected from leaf positions ranging from 5 through 22 (counting from the bottom) on infected plants and each were scanned on a flatbed scanner (Canoscan, Canon, Tokyo, Japan) 28 days after inoculation. Total pustule surface area for each leaf was determined using the ASSESS software package (version 2.0, American Phytopathological Society, St. Paul, MN). Leaves then were mounted to the inside surface of 60-mm-diameter plastic Petri dish lids with petroleum jelly. Following a modification of a technique used by Firman and Martin (8), mounted leaves were soaked 5 min. with Tween 20 water (1 drop Tween 20 per 100 ml water) (Sigma-Aldrich Co., St. Louis, MO) and blotted dry. Soaking with Tween water improved sporulation of the pathogen. Lids were placed on Petri dish bottoms containing 7 ml of a 0.05% agar solution per dish. The inclusion of a very low concentration of agar kept the released basidiospores, which dropped from leaves into the solution, suspended and thus not adhering to the surface of the Petri dish. This facilitated accurate quantification of basidiospores liberated from germinating teliospores. Dishes were placed in a plastic box, which was incubated in darkness in a growth chamber at 17°C. Two days later, the number of basidiospores deposited into the agar solution in each dish was determined by removing 10 microliters of suspended spores and quantifying with the aid of a hemocytometer. Based on the mean number of basidiospores for three replicate counts per dish and total pustule surface area per leaf, the number of basidiospores produced per mm²-pustule surface area was determined for each leaf. Comparison of replicate counts indicated that basidiospores following agitation of the collection solution were well distributed.

Lesion density varied from light to heavy regardless of leaf position on plants. A comparison of results for individual leaves indicated no relationship between numbers of collected basidiospores per unit pustule surface area with either leaf position or lesion density (data not shown). Whereas some leaves with dense lesions sporulated heavily, others with a similar lesion density had little or no sporulation. The large variation in basidiospore production later was verified in numerous experiments in which sporulation was quantified.

Teliospore Longevity in Infected Chrysanthemum Leaves

Longevity of teliospores imbedded in infected leaves was measured by the duration they were capable of producing basidiospores. Twenty cv. Vicki chrysanthemum plants were grown for 38 days after transplanting cuttings, and then inoculated as described above with Connecticut isolate CT11-3. After 24 h, plants were transferred to a greenhouse bench at 22 to 26°C and monitored for disease development. Over the next 63 days, five selected leaves on inoculated plants each were photographed on a weekly schedule for later determination of symptoms and pustule development (Fig. 1).

Pustules first emerged 7 and 14 days after infection (DAI) and appeared light in color. As they aged, their color became darker. Heavily infected leaves folded lengthwise as early as 14 DAI, and curled as early as 21 DAI. They became necrotic by 42 DAI. Leaves with fewer lesions generally survived longer without dropping from plants, but did develop necrotic areas adjacent to the pustules by 35 to 42 DAI (Fig. 1).

In the same experiment, at weekly intervals beginning 14 DAI 25 arbitrarily selected leaves displaying CWR symptoms were excised per time interval from the 20 inoculated plants. Each was scanned to determine total pustule area per leaf, incubated over water agar at 17°C, and number of basidiospores produced per mm²-pustule surface area determined as described above (Fig. 1). The experiment was repeated twice, once with isolate NY11-1 from New York and once with isolate PA11-7 from Pennsylvania.


Fig. 1. (A-E) Development of symptoms and pustules of the rust pathogen Puccinia horiana on the surface of an infected chrysanthemum leaf: (A) 14 DAI (days after inoculation); (B) 21 DAI; (C) 28 DAI; (D) 35 DAI; and (E) 42 DAI. Note that at 14 DAI, whitish areas have developed where pustules will later develop. By 42 DAI necrosis is apparent in the vicinity of the pustules. Pustules appear brown and not white because they have not sporulated. (F) Petri dishes set up as moist chambers for leaf assay. Note the basidiospores floating on the surface of the agar solution.


Sporulation on plants inoculated with each of the three isolates began by 14 DAI (Table 1). Because of unequal numbers of pustules among isolates and sampling dates, numbers of basidiospores collected from individual leaves varied to the extent that there were no clear trends. Although isolate PA11-7 sporulated the most at 14 days, and then decreased to a level that remained constant for the remainder of the basidiospore collection dates, the other isolates produced numbers of basidiospores that did not vary statistically among dates (Table 1). From visual observation of infected leaves in combination with sporulation data, it was evident that sporulation of all isolates ceased upon leaf necrosis and desiccation. In other experiments, teliospores in infected, but living, leaves were capable of sporulating for at least 112 days after inoculation (data not shown), but sporulation capability ceased upon leaf necrosis and desiccation. It was not possible to assess differences among isolates because each isolate was tested only once.

Table 1. Numbers of basidiospores produced per leaf, pustule surface areas (mm²) per leaf, and numbers of basidiospores produced per mm²-pustule surface areax.

Number of basidiospores
×10³ per leaf
Pustule area
Number of basidiospores ×10³
per pustule area
Isolate Isolate Isolate
14 420 cy 84 a 505 ab 75 d 53 b 72 b 5.5 a 2.0 a 9.9 a
21 1159 ab 515 a 880 a 316 a 179 a 343 a 4.4 a 4.0 a 3.0 b
28 951 abc 637 a 317 ab 333 ab 134 ab 314 a 4.2 a 5.8 a 1.6 b
35 1177 a 211 a 354 ab 368 a 123 ab 175 a 3.6 a 3.4 a 3.9 b
42 628 abc 302 a 93 b 234 bc 84 ab 88 b 3.3 a 4.3 a 2.4 b
49 731 abc z 78 b 191 c 73 b 4.5 a 2.4 b
56 605 bc 162 c 4.0 a
63 428 c 210 c 3.5 a

 x Chrysanthemum plants were inoculated with one of three pathogen isolates from Connecticut, New York, and Pennsylvania. On each date, 25 arbitrarily selected leaves for each isolate were incubated over a 0.05% (liquid) water agar solution at 17°C in the dark for 2 days. Basidiospores deposited into the collection solutions were counted and expressed as basidiospores per mm2-pustule surface area.

 y Data in each column with the same letters are not significantly different based on analysis of variance (P < 0.05).

 z No data were collected at these time periods because of a lack of infected leaves.

In another experiment, 12 to 16 infected plants were incubated in a mist tent for 24 h. Then four green and four necrotic leaves with pustules were excised from the plants and suspended over 1% (solid) agar in 5-cm-diameter Petri dishes and incubated 2 days in the dark at 17°C. Following incubation, basidiospore production was rated as "heavy" when visible to the unaided eye, "moderate" when easily seen by means of a dissection microscope, or "light" when visible at 100× magnification on a binocular microscope and not at lower magnification. The rating of "none" was used when basidiospores could not be detected. Collection of basidiospores on the solid water agar surface was deemed more sensitive than on liquid medium, but not quantitative. The experiment was conducted three times, with a total of 12 green (living) and 12 necrotic (dead) leaves. The green leaves all were rated as having produced heavy to light sporulation, but only one of 12 necrotic leaves in the three experiments produced any spores. Sporulation on that leaf was rated as light. In an ancillary experiment in which hundreds of necrotic leaves were placed in the mist tent along with susceptible Vicki plants, none of the plants became infected. We concluded that dead leaves are not a significant source of basidiospores for new infections in the field.

Inhibition of Sporulation by Light

Five chrysanthemum leaves infected with isolate PA11-7 exhibiting pustules with teliospores were cut into approximately equal halves. Each ½-leaf was attached to the inside surface of a Petri dish lid by means of petroleum jelly and soaked 5 min with Tween 20 water to increase the capacity to sporulate. Each lid with attached ½-leaf was placed on a Petri dish bottom containing 0.05% agar, and the closed dishes incubated at 17°C, either in darkness or under lights in a growth chamber. One ½-leaf from the original intact leaf received the light treatment and the other the dark treatment. After 16 h, the basidiospore concentration in each dish was determined as previously described. The experiment was repeated once with isolate NY11-1 and once with CT11-4. Results for the three experiments were combined for analyses and a summary is presented in Table 2.

Table 2. Mean number of basidiospores produced per leaf piece, mean pustule surface area per leaf piece, and mean number of basidiospores collected per mm²-pustule area for three P. horiana isolates

Isolate Light
Number of
Pustule area
Mean spores
per area
CT Dark 1,040,200         66.2 15,347       
CT Light 1,400         72.9 17       
NY Dark 1,001,000       117.2 8,214       
NY Light 12,600       114.1 65       
PA Dark 2,362,500       177.9 14,623       
PA Light <1       192.3 <1       

Analyses using ANOVA (P < 0.05) indicated that mean numbers of basidiospores produced per mm²-pustule area was significantly impacted by presence or absence of light regardless of isolate. For each isolate, a period of darkness dramatically stimulated basidiospore production and/or release compared to continuous light treatment. Isolate PA11-7 produced more basidiospores than isolate NY11-1 or CT11-4. Statistical comparisons among isolates, however, were not possible because isolates were tested in separate experiments. Analyses using ANOVA (P < 0.05) indicated that the presence or absence of light was not a factor in pustule size, as was to be expected because the ½-leaf pieces were randomly assigned after harvest to light or dark.


In order to determine the role of P. horiana teliospores in perpetuation of the pathogen, it was necessary to develop a more quantitative method to measure teliospore viability. To do that, two major obstacles had to be overcome. First was the high variability in teliospore germination among similarly infected leaves. This was overcome by using a relatively high number (i.e., 25) of infected leaves per sample date in the teliospore longevity study. (The required sample size was determined from a preliminary study in which variability was determined.) The second obstacle was the high propensity of released basidiospores to stick to surfaces making quantification difficult. This was overcome by adding a small amount of agar to thicken the water.

Using the new method, we showed that under the right environmental conditions teliospores could germinate as early as 14 days after inoculation of plants. Sporulation continued for the life of the leaves. Once they died, teliospores within infected leaves failed to germinate under otherwise optimum conditions. The inability to germinate was considered an indication teliospores were dead. This is the first report of the extreme susceptibility of P. horiana teliospores to leaf necrosis. This might seem contradictory to observations of Kim et al. (12) and O’Keefe and Davis (13) in which they reported overwintering of P. horiana in the field. However, whereas we measured longevity of teliospores imbedded in infected chrysanthemum leaves, it is possible they were observing survival of the pathogen through winter in systemically infected plants.

Light was shown to inhibit basidiospore production and/or spore release. Regardless of the specifics of the inhibition, results from the study suggest that it might be possible under greenhouse conditions to prevent CWR epidemics by controlling lighting. The goal would be to interrupt pathogen development without adversely affecting the chrysanthemum crop.

A better understanding of the life cycle of P. horiana, particularly where it pertains to overwintering in the field, provides potential for controlling CWR. Information gained will be of benefit for making important regulatory decisions. Results from this study show that it is highly unlikely infection of plants in the field following winter is the result of teliospores.


Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the United States Department of Agriculture. USDA is an equal opportunity provider and employer.


This research was made possible, in part, by a Cooperative Agreement from the United States Department of Agriculture’s Animal and Plant Health Inspection Service (APHIS). It may not express APHIS’ views.

Literature Cited

1. Baker, J. J. 1967. Chrysanthemum white rust in England and Wales 1963-66. Plant Pathol. 16:162-166.

2. Bonde, M. R., Peterson, G. L., Rizvi, S. A., and Smilanick, J. L. 1995. Myclobutanil as a curative agent for chrysanthemum white rust. Plant Dis. 79:500-505.

3. California Department of Food and Agriculture. 1991. Plant pathology highlights. Calif. Plant Pest Dis. Rep. (Oct.-Dec.) 10:87-89.

4. California Department of Food and Agriculture. 1992. Plant pathology highlights. Calif. Plant Pest Dis. Rep. (Jan.-May) 11:18-19.

5. California Department of Food and Agriculture. 1992. Plant pathology highlights. Calif. Plant Pest Dis. Rep. (June-Sept.) 11:36.

6. CMI. 1989. Distribution Maps of Plant Diseases No. 403, 4th Edn. CAB Intl., Wallingford, UK.

7. Exley, P. J., Giles, R. J., Pascoe, I. G., and Guy, G. L. 1993. The impact and control of white rust of chrysanthemums in Australia. (Abstr.) 6th Intl. Congr. Plant Pathol. Palais Des Congrès de Montréal: July 28-August 6, 1993, Montréal, Canada. National Research Council, Ottawa, Canada.

8. Firman, I. D., and Martin, P. H. 1968. White rust of chrysanthemums. Ann. Appl. Biol. 62:429-442.

9. Griesbach, J. A., Milbrath, G. M., and Thomson, T. W. 1991. First occurrence of chrysanthemum white rust caused by Puccinia horiana on florists’ chrysanthemum in Oregon. Plant Dis. 75:431.

10. Hennings, P. 1901. Einige neue japanische Uredineen. Hadwigia 40:25-26.

11. Hiratsuka, N. 1956. Three species of chrysanthemum-rusts in Japan and its neighboring districts. Sydowia, Ser. 2. Suppl. 1:33-34.

12. Kim, S. H., Olson, T. N., Nikolaeva, E. V., and Kang, S. 2011. Overwintering of chrysanthemum white rust caused by Puccinia horiana in Pennsylvania and challenges in its management. Phytopathology (Suppl.) 101:S91.

13. O’Keefe, G., and Davis, D. D. 2012. First confirmed report that Puccinia horiana, causal agent of chrysanthemum white rust, can overwinter in Pennsylvania. Online. Plant Dis.

14. Peterson, J. L., Davis, S. H., Jr., and Weber, P. V. V. 1978. The occurrence of Puccinia horiana on chrysanthemum in New Jersey. Plant Dis. Rep. 62:357-360.