Method To Immobilize the Aphid-Pathogenic Fungus Erynia neoaphidis in an Alginate Matrix for Biocontrol

Abstract

Erynia neoaphidis is an important fungal pathogen of aphid pests worldwide. There have been few reported attempts to formulate this natural agent for use in biocontrol. In the current study, factors involved in the immobilization of E. neoaphidis hyphae in an alginate matrix were investigated. Hyphae of two isolates cultured in liquid medium were 220 to 620 μm in length and 7 to 19 μm in diameter with a 74 to 83% cytoplasmic content. The optimal concentration of low-viscosity sodium alginate for production of conidia from entrapped hyphae was 1.5% (wt/vol), and 0.1 and 0.25 M calcium chloride were equally suitable for use as the gelling solution. Alginate beads were rinsed with 10% sucrose after gelling. However, beads should not be left for longer than 40 min in 0.1 M calcium chloride or 10% sucrose to prevent a 10% loss in conidial production. A 40% (vol/vol) concentration of fungal biomass produced significantly more conidia than either 20% or the standard concentration of 10%. This effect persisted even after beads were dried overnight in a laminar flow hood and stored at 4°C for 4 days. Conidia from freshly produced alginate beads caused 27 to 32% infection in Pea aphids as determined by standardized laboratory bioassays. This finding was not significantly different from infections in aphids inoculated with fresh mycelial mats or plugs from Petri dish cultures. In conclusion, algination appears to be a promising technique for utilizing E. neoaphidis in the biocontrol of aphid pests.

Erynia neoaphidis Remaudière and Hennebert (Zygomycetes: Entomophthorales) is one of the most widely distributed fungal pathogens of aphids, and it is an important natural factor for reducing pest aphid numbers in many crops (28). As with other members of the Entomophthorales, the primary conidia of E. neoaphidis are actively ejected from an infected sporulating cadaver under very humid conditions. Ejected primary conidia can start another infection cycle if they land on the integument of a suitable host insect. However, if primary conidia land on unsuitable surfaces (e.g., leaf or soil), then secondary conidia may be ejected that are also infective or which can in turn form tertiary and quaternary spores (8, 29).

Use of E. neoaphidis for biocontrol by dispersing sporulating cadavers or moribund but infected aphids gave mixed results (5, 27, 30). The hyphal or mycelial stage of E. neoaphidis can easily be produced in vitro, but spray applications of unformulated hyphae did not provide adequate control in glasshouse or field tests (16, 25). One possibility for using E. neoaphidis is to encapsulate the hyphae in a suitable matrix. Algination was considered to be the most obvious method, as it involves a relatively benign gelling reaction operating at ambient temperatures and is especially suitable for temperature-sensitive cells or organisms (13, 21). Sodium alginate is a positively charged polysaccharide obtained from marine algae, and matrices are formed by cross-linking with multivalent (normally divalent) cations in an ionotropic gelation reaction (3, 13). The algination technique has been used with many other beneficial microorganisms, including biocontrol fungi (1, 6, 15, 19, 26).

The objectives of the present study were to investigate factors which could affect fungal conidiation after hyphal encapsulation and to determine whether primary conidia ejected from alginate beads were still infective against aphid hosts.

MATERIALS AND METHODS

Fungal isolates.

The two isolates of E. neoaphidis used in this study were derived from single aphid cadavers collected in 1996 in Switzerland. Isolate 102 was obtained from the Pea aphid, Acyrthosiphon pisum, while isolate 158 was obtained from the Rose aphid, Macrosiphum rosae (23). Stock cultures of these isolates are stored at −80°C in 10% glycerol at the Microbiology Institute, Eidgenössische Technische Hochschule (ETH), Zürich, Switzerland. Routine cultures of the isolates were maintained on Sabouraud-dextrose agar with egg yolk (3% Sabouraud-dextrose, 1.7% agar, 8% egg yolk) at 20 to 22°C inside humid plastic boxes (12).

Liquid cultures.

Fungal mycelia were obtained in a two-step process from liquid media. In the first, preculture step, five to eight plugs (each 5 mm in diameter) of fungus were removed from a culture growing on an egg yolk agar plate and added to 20 ml of semi-skim milk medium (1.6% glucose, 1% yeast extract, 10% milk) in a 100-ml Erlenmeyer flask. After 2 days growth on a rotary shaker at 170 rpm and 20°C, 7- to 10-ml samples were added to 40 ml of fresh semi-skim milk medium in 100-ml Erlenmeyer flasks. Cultures were maintained in this second step on a rotary shaker at 170 rpm and 20°C for a further 5 days.

Hyphal dimensions.

Hyphal characters (total length, cytoplasmic length, and diameters and numbers of sidebranches) were recorded for isolates 102 and 158 after 3, 5, or 7 days of incubation in flasks during the second liquid culture step. Measurements were performed on hyphae in slide preparations with a 0.74-mm ocular scale fitted to a light microscope at ×10 magnification. Thirty hyphae were measured for each isolate at each time period.

Encapsulation of alginate beads.

Methods used in this study for alginate encapsulation of E. neoaphidis are based on procedures previously described for other microorganisms and animal cells (7, 9, 11, 14, 18, 19, 26). Fungal mycelia from liquid cultures were filtered with a sterilized brass sieve (0.71 mm) held over a 50-ml plastic centrifuge tube. The filtrate was centrifuged (2,600 × g; Universal 16R bench centrifuge, rotor 1624) at 20°C for 8 min, and the supernatant liquid was discarded. The pellet was resuspended with a previously autoclaved solution of sodium alginate (low viscosity [250 cP]; batch no. LOT 65HO335; Sigma Chemical Co.), 0.01 M MOPS (3-morpholinepropanesulfonic acid) and 0.15 M sodium chloride. The solution had been adjusted to pH 7.0 before the autoclaving. Unless otherwise stated, a final concentration of 1.5% (wt/vol) sodium alginate and 10% (vol/vol) fungal biomass was used. The alginate-fungus mixture was added dropwise from a height of 20 to 50 mm with sterilized 10-ml glass burettes (nominal diameter, 2.0 mm) into an autoclaved solution of calcium chloride (normally 0.1 M, adjusted to pH 7.0), where beads formed instantaneously. The beads were left in this gelling solution for 5 min before being harvested by filtration through a 0.18-mm sieve, rinsed with sterilized 10% (wt/vol) sucrose solution, and used directly or dried overnight in a laminar flow hood under ambient conditions of 20 to 22°C and 30 to 40% relative humidity. For storage experiments, dried beads were kept in plastic Petri dishes at 4°C for 4 days.

Concentrations of sodium alginate, calcium chloride, and fungal hyphae.

To determine the optimal sodium alginate concentration, hyphae of isolate 158 were immersed in different amounts of sodium alginate (final concentrations of 0.7, 1.5, or 3.0% [wt/vol]) for 5 min; beads were then formed by adding the hyphae and alginate solution dropwise into 0.1 M calcium chloride. After 5 min, beads were removed and rinsed for 30 s in 10% sucrose. To determine conidial numbers, groups of 15 to 20 beads of each treatment were placed onto 1.5% distilled-water agar (DWA). For the optimal concentration of calcium chloride, hyphae were mixed with sodium alginate (1.5% [wt/vol], final concentration) and then added dropwise to either 0.05, 0.1, or 0.25 M calcium chloride. Beads were left for 5 min before being washed with 10% sucrose and placed onto 1.5% DWA. The effect of varying the amounts of fungal hyphae on conidiation was studied by adding hyphae to sodium alginate to obtain a final concentration of 1.5% (wt/vol) alginate with either 10, 20, or 40% (vol/vol) mycelium. Beads were formed in 0.1 M calcium chloride and washed with 10% sucrose before being placed onto 1.5% DWA or being dried overnight in a laminar flow hood.

Effect of immersion time in solutions used for algination.

Beads were formed with 10% (vol/vol) mycelium of E. neoaphidis isolate 158 in 1.5% alginate and were left in either 0.1 M calcium chloride or 10% sucrose after a 5-min gelling period for between 0.5 and 300 min, after which samples of 15 beads were removed and placed on 1.5% DWA to assess conidial production.

Time course for production of primary conidia.

Beads were formed by using E. neoaphidis isolate 158 (1.5% [wt/vol] alginate, 10% [vol/vol] biomass). Sets of fresh beads and beads dried overnight were placed onto 1.5% DWA, and only primary conidia were counted every second day, after which beads were transferred to fresh DWA plates and incubated at 20°C under high humidity.

Conidial assessments.

A standard method was devised to assess the effects of treatments on fungal conidiation. The numbers of discharged conidia was determined by placing 15 to 20 fresh or dried beads onto plastic Petri dishes (90 mm in diameter) containing 1.5% DWA and followed by incubation at 20°C at high humidity in plastic boxes. After 5 days of incubation, individual beads were viewed with an inverted light microscope at a ×10 magnification. Conidia (primary and higher orders) present on the agar surface within a 1.5-mm radius of each bead were counted with the aid of an ocular 1-mm² grid. Counts were made at the top, bottom, right, and left of each bead and were averaged to give a single figure for each bead. For most experiments, the total numbers of conidia (primary and higher orders) were counted. The only exception was with the experiment assessing sporulation at 2-day intervals, when only primary conidia were recorded.

Infectivity of conidia.

Experiments were performed with isolates 102 and 158 to determine whether conidia were still infective after the algination process by using bioassay procedures published elsewhere (23). Beads, mycelia extracted from shake cultures with filter paper (Whatman no. 2), or plugs from egg yolk agar culture plates were inverted over young nymphs of the Pea aphid, A. pisum. In this manner, the aphids were exposed to a continual discharge of conidia (or showering) from the various inoculum sources. Three replicate plastic cups were used for each treatment, and each cup contained 8 to 20 first- to second-instar nymphs of A. pisum. With the alginate treatment, five newly produced beads were attached to the inside of each plastic lid by using small quantities of petroleum jelly as an adhesive. With the culture plug and mycelial treatments, three disks (8 mm in diameter) were placed on the insides of the plastic lids without adhesive. After the aphids had been exposed to conidial discharges for a defined time period, they were transferred to fresh pea leaves, and primary conidia were counted on coverslips that had been placed adjacent to leaf disks inside the cups. After 6 days, the numbers of living or infected aphids were assessed.

Statistical analysis.

Repeated-measure multivariate analysis of variance (ANOVA) (22) was carried out on data when a time factor was involved, i.e., hyphal dimensions, effects of varying the fungal hyphal concentrations, and assessments of primary conidia performed every 2 days. Standard univariate ANOVA was performed on data involving concentrations of alginate, calcium chloride, and conidial doses and infectivity in bioassays. Exponential regression equations were fitted to data from beads immersed in calcium chloride or sucrose. For ANOVA, data were transformed prior to analysis (9, 24). A log₁₀(x + 1) transformation was used throughout, except for the percent cytoplasmic content of hyphae, for which a square root (x + 0.5) was used, and the percent infectivity of aphids, for which no transformations were performed. Means and standard errors (SE) presented in the text refer to detransformed values where applicable.

RESULTS

Hyphal dimensions.

The structural sizes of hyphae are an important consideration in encapsulation since homogeneous sizes and shapes are highly desirable. Slide preparations of E. neoaphidis hyphae grown in liquid culture were viewed under a light microscope and measured at various culture ages. Isolates 102 and 158 were generally 220 to 620 μm in length with a diameter of 7 to 19 μm (Table 1). Hyphae of isolate 158 were significantly longer than those of isolate 102 (P < 0.0001), with a smaller diameter (P < 0.0001) and more sidebranches per hypha (P < 0.001). Significant interactions between isolate and culture age indicated hyphal length (P < 0.01) and numbers of sidebranches (P < 0.01) decreased for isolate 158 compared to isolate 102 as the cultures became older. Hyphal diameter increased with culture age for isolate 158 but not for isolate 102 (P < 0.0001). Cytoplasm in actively growing hyphae is concentrated at the tip, and large empty cell compartments can be observed behind the apical region. There were no statistically significant differences in cytoplasmic content between the isolates (P > 0.05). In addition, there was no interaction between culture periods and isolates (P > 0.05), indicating that the hyphae from the younger cultures did not contain proportionally more cytoplasm than the hyphae from the older cultures.

TABLE 1.

Mean values for hyphal dimensions of E. neoaphidis used in algination^a

Shake culture age (days)	Isolate no.	Mean hyphal:		No. of sidebranches	% Cytoplasmic content
		Length (μm)	Diam (μm)
3	102	219.9 (30.5) b	17.9 (0.6) a	0.2 (0.1) b	83.2 (4.2) a
	158	621.6 (30.5) a	7.2 (0.6) b	1.3 (0.1) a	73.9 (4.2) a
5	102	255.9 (27.3) b	19.1 (0.6) a	0.6 (0.1) a	80.7 (4.6) a
	158	363.3 (27.3) a	15.0 (0.6) b	0.8 (0.1) a	81.9 (4.6) a
7	102	311.2 (31.2) b	19.1 (0.6) a	0.2 (0.1) a	77.1 (4.6) a
	158	471.4 (31.2) a	16.5 (0.6) b	0.2 (0.1) a	77.5 (4.6) a

^an = 30 for each isolate at each shake culture age. The standard errors of the means are given in parentheses. For each variable, the different letters indicate significant differences between the means of the two isolates at each shake culture age (Tukey’s Honestly Significantly Different test (HSD), P < 0.05). 

Concentrations of sodium alginate, calcium chloride, and fungal hyphae.

Conidial numbers discharged from alginate beads were used as the indicator for hyphal viability after different treatments. A sodium alginate concentration of 1.5% (wt/vol) produced a significantly higher number of conidia than either 0.7% or 3.0% (wt/vol) alginate (P < 0.0001), an amount almost threefold greater than the other two concentrations evaluated (Table 2). With 1.5% (wt/vol) sodium alginate, beads gelled in 0.1 or 0.25 M calcium chloride for 5 min resulted in significantly higher sporulation than beads kept in 0.05 M calcium chloride (P < 0.0001) (Table 2).

TABLE 2.

Effects of sodium alginate and calcium chloride concentrations on conidiation from freshly produced alginate beads of E. neoaphidis isolate 158

Treatment	Concn		No. of conidia mm−2a
	% (wt/vol)	M
Sodium alginate	0.7		35.0 b
	1.5		104.3 a
	3.0		36.6 b
Calcium chloride		0.05	79.1 b
		0.10	117.1 a
		0.25	118.7 a

^aThe standard error of the mean for all values was 0.1. With either the alginate or the calcium treatment, the different letters indicate significant differences between the means (Tukey’s HSD, P < 0.05). 

Increasing the amount of fungal mycelia encapsulated with sodium alginate significantly enhanced conidial production (P < 0.0001). Sporulation from fresh beads containing 40 and 20% (vol/vol) mycelia resulted in 35.3 (SE = 0.1) and 20.1 (SE = 0.1) conidia mm⁻², respectively. This result was 13- to 22-fold greater than the number from 10% (vol/vol) mycelia (1.6 ± 0.1 conidia mm⁻²). Drying the beads overnight and then storing them for 4 days caused the conidial numbers to be significantly reduced by 63 to 97% compared to fresh beads (P < 0.0001). Upon rehydration, dried beads with 40% (vol/vol) mycelia produced 5.9 (SE = 0.1) conidia mm⁻², which was 10-fold higher than the value of 0.6 (SE = 0.1) conidia mm⁻² obtained from either the 10 or 20% (vol/vol) mycelial concentration.

Conidiation after immersion in solutions used for alginate bead production.

To determine the optimal time for keeping beads in either solutions for gelling (0.1 M calcium chloride) or for rinsing (10% [wt/vol] sucrose), beads were left in either of the solutions for different periods of time at an ambient temperature. Conidial counts from fresh beads showed a decline with time for both solutions, and there was good agreement with the exponential regression equation y = ae^bx, where y is the conidial number and x is the time in solution. For immersion time in calcium chloride, values of a and b were 110.1 (SE = 1.1) and −0.0025 (SE = 0.0005), respectively (r² = 0.95, n = 5, P < 0.05). For immersion in sucrose, a and b were 116.5 (SE = 1.1) and −0.0029 (SE = 0.0006), respectively (r² = 0.81, n = 6, P < 0.05). With the derived equations, 10 and 50% reductions in conidial numbers are predicted if beads are kept in 0.1 M calcium chloride for 42 and 273 min, respectively. For 10% (wt/vol) sucrose, 10 and 50% reductions in conidial numbers are predicted after 37 and 243 min, respectively.

Production of primary conidia.

The numbers of primary conidia ejected onto 1.5% DWA from fresh and previously dried beads were monitored over a 9-day period. From five sample dates, a total of 245 primary conidia were counted from fresh beads and only 68 primary conidia were counted from dried beads. This was equivalent to a 72% reduction in conidia directly attributable to overnight drying. For both fresh- and dried-bead treatments, the conidial numbers were also affected by rehydration time. With fresh beads, the mean numbers of primary conidia varied between 0.72 and 1.09 (SE = 0.05) conidia mm⁻² on days 1, 3, and 5 and were significantly higher than on days 7 and 9 (P < 0.0001), where they declined to 0.2 and 0.02 (SE = 0.05) conidia mm⁻², respectively. With dried beads, mean values of 0.02 to 0.16 (SE = 0.04) conidia mm⁻² on days 1, 3, 7, and 9 were significantly lower than on day 5 (P < 0.0001), when they reached a maximum of 0.40 (SE = 0.04) conidia mm⁻². Hence, conidial discharge was delayed and peaked on the fifth day of rehydration for dried beads compared to fresh beads.

Infectivity of discharged conidia.

To apply sufficient numbers of primary conidia for infection, aphids were showered for 6 h with isolate 102 and overnight with isolate 158. There were significant differences between treatments in the numbers of primary conidia applied during bioassay tests with isolate 102 (P < 0.01) and isolate 158 (P < 0.001). In both cases, the doses applied with alginate beads were significantly lower than for the treatments with fresh mycelia or the plugs taken from agar cultures. However, no significant differences were detected in infection between the alginate beads and the other inoculum sources with either isolate 102 or 158 (P > 0.05) (Table 3).

TABLE 3.

Mortality in first-instar Pea aphids with formulated and unformulated E. neoaphidis^a

Isolate	Inoculum form	Conidial dose (no. of primary conidia mm−2)	% Infection	No. of aphids used
102	Extracted mycelium (mats)	284.8 (0.2) a	42.3 (2.7) a	59
	Culture dish (plugs)	207.1 (0.2) a	42.3 (2.7) a	51
	Alginate beads	65.4 (0.2) b	31.8 (2.7) a	69
158	Extracted mycelium (mats)	112.8 (0.2) a	35.8 (4.4) a	43
	Alginate beads	13.2 (0.2) b	26.5 (4.4) a	47

^aStandard errors of the means are given in parentheses. With each of the isolates, values with the different letters indicate significant differences between the means for either the conidial dose or the percent infection (Tukey’s HSD, P < 0.05). There was no infection in uninoculated controls. 

DISCUSSION

There are surprisingly few reports on the hyphal dimensions of E. neoaphidis grown in vitro. With a semidefined liquid medium, an isolate of E. neoaphidis decreased in hyphal length from 325 to 155 μm after 8 days of cultivation (10). This finding is similar to the behavior of E. neoaphidis isolate 158 in the current study. Hyphae of E. neoaphidis from infected insects are reported to be smaller than those obtained from solid or liquid medium, with lengths of 32 to 260 μm and diameters of 6 to 13 μm (10, 28). Currently, mycelia from shaking-flask cultures are harvested at 5 days. Cultures older than this tend to form mycelial pellets (ca. 2 to 3 mm in diameter), resulting in a lowered biomass for filtering and centrifugation prior to algination. Conversely, liquid cultures younger than 5 days have very low yields of biomass. Algination appears to be a useful technique for the encapsulation of a fungus such as E. neoaphidis, which grows as large, nonuniform hyphal fragments in artificial media. Even so, only filtered mycelia are used at present since unfiltered mycelia block the orifices of glass burettes used to add fungus and alginate mixture into the gelling solution. Fungal biomass could be blended with alginate (18), but this was found to be inappropriate for E. neoaphidis in preliminary experiments since the hyphae are mostly aseptate and cytoplasmic disruption occurs during maceration.

Increasing the amount of E. neoaphidis mycelium in alginate solutions from 10 to 40% (vol/vol) increased conidiation by 10- to 22-fold with dried and fresh beads, respectively. The use of 0.1 or 0.25 M calcium chloride as gelling agent is preferred to using 0.05 M. Beads should not be kept in 0.1 M calcium chloride or 10% sucrose for longer than 40 min so as to prevent a reduction in conidiation. Preferably, beads should only be kept for 1 to 5 min in either of the two solutions. There were no differences in aphid infection between the alginate beads, the culture plates, and the crude mycelial extracts; hence, the conidia remained infective after algination. One practical advantage of using alginate beads with E. neoaphidis (and possibly other Entomophthorales) is that active discharge of conidia occurs from the bead surface. Hence, infection may be less dependent on chance contact between a suitable aphid host and a sporulating bead, which is likely if the conidia are simply attached to conidiophores.

There are two factors that have not been addressed in the present study but which may affect the productivity of entrapped cells in alginate beads. First, polysaccharide components differ among alginates extracted from different seaweed sources (2), but batches of sodium alginate obtained from commercial suppliers may be purified with an organic solvent before use (20). Second, autoclaving of sodium alginate is not recommended as it reduces alginate viscosity and gel strength (17). However, this depolymerization can be ameliorated by buffering sodium alginate solutions at pH 7 to 8 (4), and autoclaving is preferable for laboratory studies as it minimizes saprophytic contamination of beads during assays.

Algination appears to be a promising technique for encapsulating hyphae of E. neoaphidis produced in vitro to be used against aphids for biocontrol. Further studies are in progress to improve the conidiation and storage of alginate beads by the addition of nutrients to alginate solutions and by using milder drying procedures.