Take-All Root Rot

A Detrimental Root Disease of Bermudagrass Putting Greens

By Maria Tomaso-Peterson, Ph.D and Aaron Tucker, MSc., Mississippi State University Cameron Stephens, Ph.D candidate, North Carolina State University

Take-all root rot (TARR), as the name indicates, is a disease that affects roots, stolons, and rhizomes. This disease is widespread on ultradwarf bermudagrass greens throughout the transition zone and subtropical climates, including Alabama. However, TARR is also active in dwarf bermudagrass greens and sod production fields where bermudagrass putting green cultivars are grown. Until recently the causal agent of TARR was identified as Gaeumannomyces graminis var. graminis – or Ggg. We initiated research in 2012 in the Turfgrass Pathology lab at Mississippi State University (MSU) to get a better understanding of the ectotrophic root-infecting (ERI) fungi associated with this disease — originally referred to as bermudagrass decline (BD). We identified three novel ERI fungi, G. nanograminis (Gn), Magnaporthiopsis cynodontis (Mc), and Candidacolonium cynodontis (Cc), that formed a complex with G. graminis infecting roots in bermudagrass (BG) putting greens. These fungi do not discriminate between Tifdwarf, 328, or other ultradwarf bermudagrasses. Parallel research conducted by Cameron Stephens and Dr. Jim Kerns at North Carolina State University (NCSU) identified a fifth ERI fungus, G. graminicola (Ggram), associated with root-rot and symptoms of TARR. ERI fungi colonize the outer surface of roots, stolons, and rhizomes and produce distinct structures, readily identifiable with the aid of a microscope. They are characterized by the formation of dark brown to black runner hyphae and growth cessation structures on the surfaces of roots, stolons, and rhizomes. Infection hyphae originate from lobed hyphopodia (Fig. 1).

FIGURE 1: Infection hyphae originate from lobed hyphopodia.

The foliar symptoms of TARR first appear as irregularshaped, chlorotic to white patches up to 3.0–ft in diameter. Foliar symptoms of chlorosis/necrosis progress upwards from the point of infection near the crown of the plant to the leaf tips. The chlorotic to white patches may be solitary or coalesce with neighboring patches to create large, irregular areas within a green. A general thinning with the patches or along the margin of the green may result if the disease is left unchecked (Figs. 2–3). These symptoms may be confused with other foliar diseases such as mini ring. Therefore, it is important to check the root system of affected plants. These roots, stolons, and rhizomes will appear dark brown to black. Black lesions may be observed on the stolons and roots. Overall, the roots will be very short, 0.5–in., brown, rotted, brittle and lack secondary roots and root hairs.

FIGURE 2: Chlorotic to white patches appear solitary or coalesce with neighboring patches.

FIGURE 3: Chlorotic to white patches appear solitary or coalesce with neighboring patches.

Take-all root rot symptoms occur during or immediately after warm to hot, humid periods in the late summer or early autumn when the day length shortens. The chlorotic patches tend to persist into the winter months when greens are covered to protect from freezing temperatures. In dormant BG greens, the remnant patches are usually obvious, becoming more conspicuous during spring green-up, and fade as vigorous BG growth resumes in the early summer. TARR symptoms are not typically observed during the summer months, but does that mean the roots are void of ERI fungal infection? In 2017–18, MSc. student at MSU, Aaron Tucker, conducted an extensive survey of ERI fungi associated with BG roots from two putting greens at the MSU golf course. One green was considered healthy or asymptomatic and the other had a history of TARR or BD. We sampled 68 areas of interest (AOI) from the healthy green and 66 AOIs from the TARR green. The roots were collected from annual July core aerification events in both years. A rapid molecular assay was developed specifically to identify the four ERI fungi of interest, Gg, Gn, Mc, and Cc. In general, at least one ERI fungus was identified in each AOI from both greens. Gn had a high frequency of occurrence (FO) in the healthy green and Gn had a higher FO in the TARR green. Mc was identified in the roots of nearly all AOIs from both greens and Cc was identified in approximately 50% of AOIs in both greens each year. We also observed the ERI fungi formed a complex at a high frequency within sampled roots. In the healthy green in 2017, 7% AOIs had a fungal complex composed of the four ERI fungi and in 2018 50% AOIs had a Gn, Cc, and Mc complex. Multiple complex variations were observed in the TARR green in both years. The four ERI fungal complex was observed in 7.5% and 15% AOIs for 2017 and 2018, respectively. A three ERI fungal complex consisting of Gn, Mc, and Cc was observed in 27% AOIs in 2018. Previous research at MSU and most recently at NCSU showed the individual ERI fungi express variable levels of aggressiveness on bermudagrass roots. This indicates each ERI fungus alone can cause root rot. They also have different levels of fungicide sensitivity, growth rates, optimal temperatures.

In vitro fungicide sensitivity assays to screen the ERI fungi were conducted independently by Tucker and Stephens. Both showed the ERI fungi are not sensitive to succinate dehydrogenase inhibitor (SDHI) fungicides such as fluxapyroxad, penthiopyrad, and isofetamid. The ERI fungi showed moderate levels of sensitivity to the strobilurin (QoI) and demethylation inhibitor (DMI) fungicides compared to the SDHIs with azoxystrobin and pyraclostrobin having the greatest inhibitory effect on fungal colony growth. Tucker’s results at MSU calculated the effective concentration to inhibit 50% (EC 50 ) colony growth. No fungicide completely inhibited fungal growth (Table 1). How do these results transfer to fungicide control in a BG putting grass?

Take-all root rot pathogen growth was evaluated at 50, 59, 68, 77, 86, and 95°F to determine optimal growth temperature of each organism. In general, all TARR pathogens grew optimally between 77 and 86°F (Fig. 4). Optimal growth temperature for Gg, Gx, Ggram, Cc, and Mc was 78.4, 75.4, 76.3, 91.0, and 80.1°F, respectively. Historically, TARR has primarily been viewed as a fall season issue with applications for TARR management typically beginning in late September-October. However, results from this study indicate these pathogens are growing optimally and potentially causing the most damage to bermudagrass roots through the summer months. That damage may manifest in the fall and spring as characteristic TARR symptoms when adverse bermudagrass growing conditions such as low light and cooler temperatures are present. By that time, the damage has been done, the pathogens are not actively growing, and curative fungicide applications may be futile. Ultimately, this information warrants earlier fungicide applications than traditionally applied. For fungal growth to be inhibited, the fungus must be actively growing to absorb the fungicide. Corresponding the optimal growth temperature information from this study to the soil temperature data at a 2-4-in. depth from Starkville, MS (Fig. 5), we can see TARR pathogens may be growing optimally from May-Sept in 2019 and June-Sept in 2020. Ideally, this is when we should be applying fungicides for TARR management.

FIGURE 4: Growth of Gaeumannomyces graminis (Gg), Gaeumannomyces nanograminis (Gx), Gaeumannomyces graminicola (Ggram), Candidacolonium cynodontis(Cc), and Magnaporthiopsis cynodontis (Mc) at different incubationtemperatures. Means followed by the same letter within temperature arenot statistically different according to Fisher’s Protected LSD test atP<0.05.

FIGURE 5: Average soil temperatures of the top 4.0 inches in Starkville, MS. The red bars indicate the optimal temperature range for ERI fungal growth.

Fungicide application timing research has also been conducted at NC State to validate the previous claims under field conditions. Field studies were initiated evaluating Tartan application timing for TARR management. Plots received four applications of Tartan on a 21-day interval with staggered start dates. All applications were watered in with 1/8th inch post-application irrigation. Application start dates were Timing 1 (15 July), Timing 2 (5 August), Timing 3 (26 August), Timing 4 (16 September), and a non-treated control (NTC). On 6 Jan 2020, Tartan applications initiated on Timing 2 (5 August 2019) provided the best TARR control and Timing 3 (26 August 2019) provided moderate disease suppression (Fig. 6). However, if we look at the area under the disease progress curve, or disease progression over time in 2020, Timing 1, Timing 2, and Timing 3 all significantly reduced TARR severity compared to the non-treated control (Fig. 7). Tartan applied on Timing 4 did not reduce TARR compared to the nontreated control. Timing 4 was most likely applied too late when the pathogens were not actively growing.

FIGURE 6: Take-all root rot severity on 6 January 2020 influenced by various Tartan application initiation dates. Timing 1=15 July, Timing 2=5August, Timing 3=26 August, Timing 4=16 September, NTC=Nontreatedcontrol. Means followed by the same letter are not statisticallydifferent according to Fisher’s Protected LSD test at P<0.05.

FIGURE 7: Take-all root rot severity over time (area under disease progress curve; AUDPC) influenced by various Tartan application initiation dates. Timing 1=15 July, Timing 2=5 August, Timing 3=26 August, Timing 4=16 September, NTC=Non-treated control. Means followed by the same letter for AUDPC values are not statistically different according to Fisher’s Protected LSD test at P<0.05.

A similar study was conducted evaluating Maxtima and Navicon application initiation timing for TARR management. Plots received two applications of either Maxtima or Navicon on a 21-day interval with staggered start dates. All applications were watered in with 1/8th inch of post-application irrigation. Treatments (initiation date) were Maxtima 1 (25 August), Maxtima 2 (25 October), Navicon 1 (25 August), Navicon 2 (25 October), and a non-treated control (NTC). Similar to the Tartan application timing results, the earlier applications of either Maxtima or Navicon provided the best TARR suppression on 8 October 2020, 29 October 2020, and 19 November 2020 (Fig. 8). Disease progression over time was the lowest for the Maxtima 1 and Navicon 1 application timings (Fig. 9) suggesting the earlier application start dates provided the best TARR suppression.

FIGURE 8: Take-all root rot severity influenced by various Maxtima and Navicon application initiation dates. Maxtima 1=25 August, Maxtima 2=25 October, Navicon 1=25 August, Navicon 2=25 October, NTC=Non-treated control. Means followed by the same letter within rating date values are not statistically different according to Fisher’s Protected LSD test at P<0.05.

In general, initiating fungicide applications for TARR earlier than traditionally applied may provide greater TARR control. The earlier application timings of Tartan, Maxtima, and Navicon provided greater control of TARR compared to the later application timings and the non-treated control in Raleigh, NC. Field trial results presented here align well with the optimum in vitro growth temperature study. In Central MS, targeting TARR application initiation for late May-mid-June may provide the greatest disease control. However, soil temperatures should be monitored in your area to make the most informed decision. •

FIGURE 9: Take-all root rot severity over time (area under disease progress curve; AUDPC) influenced by various Maxtima and Navicon application initiation dates. Maxtima 1=25 August, Maxtima 2=25 October, Navicon 1=25 August, Navicon 2=25 October, NTC=Non-treated control. Means followed by the same letter for AUDPC values are not statistically different according to Fisher’s Protected LSD test at P<0.05.