Burned and unburned oil-contaminated media impede lateral growth of Neurospora crassa regardless of hydrophobin expression

Julia A. DeFeo and William J. Myers
Department of Biology, Rutgers University, Camden, NJ 08102


Bioremediation, or the use of organisms to degrade environmental pollutants, is an increasingly prevalent concern. Due to surging global oil production and the correlating increase of improper burned and unburned oil disposal, more information about organisms capable of bioremediation is needed. Fungi are commonly considered for bioremediation due to their production of various exoenzymes and documented capacity for degrading crude oils. Hydrophobins, an amphipathic protein utilized by some filamentous fungi to decrease the surface tension of surrounding liquid media, may be an important consideration in the growth of fungi in the context of bioremediation. This study aims to characterize the growth of Neuropsora crassa in burned and unburned oil-contaminated media with and without the production of hydrophobins in order to determine if N. crassa can tolerate an oil-polluted environment. Here we show that both burned and unburned oil-contaminated media lead to a significant impairment of lateral growth. There was no significant impact on lateral growth due to the presence of hydrophobins. This experiment therefore demonstrates an inihibted capacity for N. crassa to grow in oil-contaminated environments. Future research should aim to further define specific growth characteristics of N. crassa in oil-contaminated environments such as effects on reproductive structures and hyphal branching.



Oil is a potent pollutant, and the threats of oil spills and improper oil disposal have increased alongside oil production. Crude oil spills can contaminate and cause massive harm to large ecosystems. Burned motor oil, which contains heavy metals and aromatic hydrocarbons from the burning process, is often not disposed of in the proper facilities, and thus can be toxic for organisms that are exposed to the oil (Akintunde et al., 2012). Better technologies need to be developed for oil spill prevention, containment, and the remediation of oil-contaminated environments.
Bioremediation efforts, or the use of organisms to degrade pollutants, encounter a range of issues based on the degree of pollution, which microorganisms are used, and what byproducts are produced (Ekundayo, F. O. et al., 2012). The amount of pollutants present can lead to adverse growth effects even on organisms that can degrade the pollutants. Determining if the chosen organism can tolerate the severity of the given pollutant is an important consideration in its capacity for bioremediation. More information on the pollution tolerance of bioremediative microorganisms is therefore needed.
Fungi are commonly considered for bioremediation because they possess a wide range of enzymes that can metabolize various highly recalcitrant substrates. Fungi are also known to be very tolerant of growth conditions that are adverse to other eukaryotes, but the exact toxicity of the contaminants in burned oil on fungi is not well-documented.
However, many fungal species have been reported to degrade crude oil (Ekundayo, F. O. et al., 2012). Fungi are therefore desirable candidates for the bioremediation of burned oil spills, due to their tolerance for toxic conditions and their oil-degrading capabilities. Further  experimentation will determine the viability of specific fungal species for oil remediation.
Neurospora crassa is a filamentous fungi that has been isolated from oil-contaminated environments and has a documented ability to degrade crude oil, as well as phenolic compounds commonly found in oils, but its efficiency with regard to degrading crude oil is generally less than that of other similarly capable fungal species (Ekundayo, F. O. et al., 2012;  Luke  et  al.,  2001). However, N. crassa does possess certain advantages that could make it a desirable selection for bioremediation. Not only is N. crassa relatively nonpathogenic, but it is also easily available and its genome is entirely mapped due to its staple use in genetic research. It is therefore feasible to genetically engineer strains of N. crassa so that they are adapted to best perform in specific ecosystems for particular bioremediation needs. This genetic consideration in particular makes N. crassa an excellent model organism for studying the possibility of genetically adapting microorganisms for specific bioremediation contexts.
One genetic consideration is the production of amphipathic proteins known as “hydrophobins” (Bayry et al., 2012). Being amphipathic proteins, they are known to self-assemble into sheets with a hydrophilic face and a hydrophobic face. This allows them to adhere to surfaces, changing their affinity for water. In fungi, hydrophobins coat spores to make them difficult to “wet,” or become saturated with water. N. crassa produces the hydrophobin EAS, which is a Type I hydrophobin (Bayry et al., 2012). Hydrophobins such as EAS are also known to reduce the surface tension of liquid (Bayry et al., 2012) and form assemblies at the interfaces of oil and water (Linder  et  al.,  2005). Hydrophobins thereby affect fungal growth by reducing the surface tension of the surrounding liquid medium so that the fungi can push its sporangia and hyphae through the fluid. This study is thus concerned with the impact the presence of hydrophobins will have on hyphal growth in an oil-contaminated medium.
The scientific record still needs to address the effect of an oil-contaminated environment on the growth of filamentous fungi. N. crassa’s tolerance for burned and unburned oil, and the nature of its growth through these contaminants, needs to be characterized. The production of hydrophobins is an important factor for fungal growth through any liquid medium, but the impact of their presence on fungal growth in burned and unburned oil-contaminated media is unknown. For the purpose of informing the bioremediative capabilities of N. crassa, this study seeks to demonstrate the capability of N. crassa to grow in an oil-contaminated environment. By exposing N. crassa to an oil-contaminated media, it will be determined if there is increased growth through the medium when the hydrophobin gene is expressed. This is expected because oil has a lower surface tension than water and hydrophobins will act to further lower this minimal surface tension. It is also predicted that there will be decreased growth in burned oil treatments, possibly due to the presence of toxic heavy metals and aromatic hydrocarbons. Therefore, the hypothesis of this study is that N. crassa producing hydrophobins in unburned oil-contaminated media will exhibit the most growth, while N. crassa not producing hydrophobins in burned oil-contaminated media will exhibit the least amount of growth.


Materials & Methods

This study used 36 replicates evenly distributed across the following treatments: Wild Type –Distilled Water; Wild Type –Burned Oil; Wild Type –Unburned Oil; Mutant –Distilled Water;  Mutant –Burned  Oil;  Mutant –Unburned  Oil.  (6 replicates per treatment)
Strains and Suspensions
Two strains are used in this study: FGSC 2489 (wild-type) and FGSC 13319 (mutant lacking hydrophobin production; also called easily-wettable (eas) or clock-controlled-gene-2 (ccg-2). These strains were provided by Dr. Kwangwon Lee.
A  conidial  suspension  was  made  for  each N. crassa strain using 48-hour old cultures. Sterilized water was pipetted into the  culture  tubes,  and  the  tubes  were  then  vortexed.  The resulting   solution   was   poured   through   autoclaved cheesecloth,   isolating   the   conidia, into autoclaved microcentrifuge tubes. Using a BioRad TC20™ Automated Cell Counter, the concentration of conidia in each suspension was determined. The suspensions were then diluted to 2.50 x 106conidia/mL.
3 L of Vogel Medium N Agar was made using 60 mL of 50X Vogel Salts, 2.94L of deionized water, 45 g of sucrose, and 45 g of agar, mixed and heated to 90 ºC and stirred at 350 rpm for 20 minutes. The agar mixture was then autoclaved. After autoclaving, 150 mL of the Vogel Agar was poured into each of 18, 12” x 3” containers. Agar was left to settle and cool for 30 minutes.

Figure  1.  Experimental  Design.    Each  large  rectangle  with  thick outline  represents  a  container.    Within  each  container  are  two medium rectangles representing treated filter paper (light blue for distilled water, peach for unburned oil, brown for burned oil) that meet and slightly overlap in the middle of the container. At either end  of  each  container  are  small  rectangles  representing  the uncovered Vogel Agar that serves as the inoculation site.  The top row  of  containers  was  inoculated  with  wild-type  FGSC  2489 N. crassa and the bottom row was inoculated with mutant FGSC 13319 N. crassa.


Containers (large rectangles depicted in Fig. 1) were assigned their treatments. Half of the 18 containers were inoculated with wild type N. crassa FGSC 2489, and the other half were inoculated with FGSC 13319. Within each of these halves, 3 containers were assigned to a distilled water treatment, 3 were assigned to an unburned oil treatment, and 3 were assigned to a burned oil treatment, which was a mixture of unburned and burned oil in a 50:50 by volume ratio (the burned oil was diluted due to a lack of results in burned oil replicates in unreported pilot studies). There were two replicates within each individual container, for a total of 36 replicates, 6 within each treatment.

Filter Paper Treatment

For each container, two 7.62 cm x 12.65 cm filter papers (medium rectangles depicted in Fig. 1)  were soaked with 1.5 mL of the assigned treatment. A micropipette was used to pipet 100 μL in a 5 x 3 grid pattern. Each 100 μL increment was separated by 2.54 cm (1.27 cm from edge of filter paper), with three increments across the width (7.62 cm) of the filter paper and five down its length (12.65 cm). Two filter papers were placed in each container so that the filter papers only slightly overlap in the middle. Each edge of the container is then left with 7.62 cm x 2.54 cm uncovered Vogel Agar (small rectangles depicted in Fig. 1), which served as the inoculation sites. This gridding pattern ensures that the filter papers are saturated and not oversaturated and that the amount of treatment is normalized. Unburned oil was Pennzoil 550035091 Motor Oil Lubricant. Burned oil was taken from a car after an oil change.


To inoculate, 100 μL of suspension (2.50 x 105conidia) was pipetted directly onto the middle of the edge where the 7.62 cm x 2.54 cm uncovered Vogel Agar and side of the container meet. Suspensions were vortexed between each inoculation.

Conditions and Measurements

The containers were stored in incubators set to 25 ºC, covered with parafilm. Parafilm was sterilized with 91% isopropyl alcohol. Growth was recorded and documented every 24 hours by measuring, with a ruler, the lateral appearance of macroscopic mycelial growth (in cm), relative to the edge of the filter paper nearest the inoculation site. Pictures were also taken of each individual replicate.



The lateral growth was measured at 24 hour intervals, but only the data from the 72 hour interval was analyzed. Pictures were also taken every 24 hours. 72 hours after inoculation, there was no statistical difference between lateral growth across both strains of N. crassa (mutant (without  hydrophobins) and wild type (with hydrophobins))  (two-way  ANOVA,  p  >  0.05)  (Fig.  2). Additionally, there is a difference between lateral appearance of mycelial growth across oil treatments (burned/unburned/control)  (two-way  ANOVA,  p  <  0.05) (Fig. 2). There was no interaction between oil treatment and fungal strain (two-way ANOVA, p > 0.05) (Fig. 2).


Figure 2. Lateral growth of N. crassa 72 hours after inoculation. Significant difference between distilled water (control) and either oil-based treatment. n = 6 for each treatment. Wild-type FGSC 2489 represented by black bars and mutant FGSC 13319 represented by white bars.

There is also no significant difference between Mutant Burned growth and Wild Type Burned growth (Tukey HSD p > 0.05), and Mutant Unburned growth was not significantly different from Wild Type Unburned growth (Tukey HSD p > 0.05) (Fig. 2).

Average Mutant Unburned was not significantly different from Mutant Burned growth (TukeyHSD p > 0.05) (Fig. 2). Average Wild Type Unburned growth was also not significantly different from Wild Type Burned growth (TukeyHSD p > 0.05) (Fig 2.).

Compared to the control replicates, which were grown in distilled water, both Mutant Burned and Mutant Unburned had significantly less growth (TukeyHSD p < 0.05) (Fig. 2). The same is true for Wild Type Burned and Wild Type Unburned compared to distilled water, each having significantly less growth (TukeyHSD p < 0.05) (Fig. 2). There was no significant difference between Mutant Control and Wild Type Control (p > 0.05) (Fig. 2).

In unburned treatments across both strains, the unburned oil spilled from the filter papers into the inoculation site. In burned oil treatments, there appeared to be localized concentrations of burned oil at the margin of hyphal growth (best seen in Figure 3, middle image). Burned oil treatments appeared to present with more fuzzy white macroscopic growth than the unburned oil treatments (Fig. 3).

Figure 3. Growth phenotypes. Images of wild-type FGSC 2489 in distilled water (A), burned oil (B), and unburned oil (C), as well as images of hydrophobin mutant FGSC 13319 in distilled water (D), burned oil (E), and unburned oil (F).



Based on the results of this trial, burned and unburned oil significantly decreased the lateral growth of the mycelium and hydrophobin presence did not have an effect in any oil treatment. There was no significant difference in lateral growth between burned and unburned oil treatments.

Mutant growth was greater than wild type growth in unburned oil, but not significantly so. However, this is unlikely to be due to an actual effect of the lack of hydrophobins, as this difference mostly appears to be the result of a single high-growth outlier in the mutant unburned treatment. This outlier resulted in the relatively high error bars for the mutant unburned treatment. More replication is necessary to determine the validity of this outlier. Higher mutant growth in any oil treatment relative to wild type growth would also not be expected because the presence of hydrophobins serves to decrease surface tension at oil-water interfaces (Linder et al., 2005). So increased lateral growth in the absence of hydrophobins would contradict the current understanding of hydrophobins.

The interaction between the hydrophobic unburned oil and the water-based agar also led to a “flooding” effect that could have confounded the results in the unburned oil treatments.  This could have affected the results as the inoculations would have immediately encountered the oil treatment, which has been shown in this study to impair growth. The burned oil treatments, which did not experience this “flooding,” did not encounter their treatment until they reached the filter paper.

The lack of a significant effect of hydrophobins on lateral growth was surprising, and so the strains were re-tested for “wettability” after the conclusion of the experiment to ensure that there had been no cross-contamination. There is no clear explanation for this lack of effect, and future studies should attempt to further characterize the role and impact of hydrophobins at hyphal tips exposed to oil.

Although there is no significant difference between unburned and burned oil, it is difficult to draw definitive conclusions about the effect of toxic compounds in the burned oil and their effect on the lateral growth. Firstly, the burned oil was diluted with unburned oil.  The  “flooding”  effect in the unburned oil treatments must also be considered. Qualitative observations support this explanation, as the inoculation sites at 24 hours appeared to be covered with a film of unburned oil, but this “flooding”  was not observed  in the burned oil treatments. There also appeared to be a “pooling” of the dark pigment characteristic of the burned oil at the margin of hyphal growth in the burned oil replicates. It may be possible that the hyphae are somehow translocating the toxins concentrated in the burned oil that they are not capable of immediately degrading.

The significantly impaired lateral growth in the oil treatments, compared to the distilled water control treatments, could also be due to nutrient richness rather than only environmental impairment. The presence of massive hydrocarbons in these treatments could impede lateral growth as the fungi might prefer to grow a more heavily-branched mycelium when it encounters these large hydrocarbons in addition to the Vogel Agar. This could be a feasible explanation due to N. crassa having a relatively limited capacity for degrading crude oil (Ekundayo, F. O. et al., 2012). Although N. crassa can degrade crude oil, it may be that it cannot degrade crude oil to a degree that would be overall beneficial for its growth.

There is also an obvious effect on the composition of the mycelium due to burned oil exposure as observed by the dominance of the fuzzy white mycelial structures in these treatments. This change in appearance could possibly be related to the reproductive structures produced along the mycelium, and consequently this difference could have an effect on mycelial growth.

Future studies should attempt to observe and quantify the branching patterns when N. crassa is exposed to burned and unburned oil. Quantifying the relative amounts of the different reproductive structures of N. crassa when exposed to oil treatments could also be of interest. Higher replication and better control of the localization of the oil treatments in future trials would lead to more definitive results. Enzyme assays could be used to characterize the biochemical mechanism by which N. crassa breaks down the compounds in the oil treatments. A future study could then also attempt to increase the expression of relevant enzymes using genetic modification.

Any replicated trials of this experimental design should also seek to use a more concentrated burned oil solution. Here the burned oil was diluted to a 50:50 by volume ratio with unburned oil. This was done because pilot studies had found that the concentrated burned oil solution led to almost no growth of N. crassa. Future trials should attempt an 80:20 or 70:30 by volume ratio of burned oil to unburned oil.

Future research should also aim to further inform the role N. crassa could play in bioremediation. There must be a clear understanding of what byproducts are formed when introducing a bioremediative microorganism into a polluted environment, and so identifying the compounds produced as the result of N. crassa degradation of oil is important. Consideration must also be given to the other effects, independent of pollution, that a specific microorganism would have on the other populations and the abiotic environment composing the ecosystem into which it is introduced. Experiments growing N. crassa in an oil-contaminated environment that is also inhabited by other fungal species would be an example of a useful study for informing this concern.



We thank Dr. Kwangwon Lee for giving us the opportunity to carry out this project through his Principles and Practices of Biological Research (PPBR) course, as well as his generous gifts of Vogel Salts, N. crassa strains, culture tubes, and cell-counting cassettes.

We thank Dr. Nathan Fried for his role in teaching the PPBR course, as well as his assistance in generating ideas for our project, analyzing  data, and providing feedback for our writing process.

We especially thank Ms. Sarah Johnson for answering all of our technical inquiries and for providing us with the materials necessary to make this project happen.

We thank Dr. Katie Malcolm and Dr. Jennifer Oberle for their helpful input based on their expertise with fungi.



Akintunde, W.O., Olugbenga, O.A., and Olufemi, O.O.(2015).  Some Adverse Effects of Used Engine Oil (Common Waste Pollutant) On Reproduction of Male Sprague Dawley Rats.  Open Access Maced JMed Sci 3, 46–51.

Bayry, J., Aimanianda, V., Guijarro, J.I., Sunde, M., and Latgé, J.-P. (2012).  Hydrophobins—Unique Fungal Proteins.  PLOS Pathogens 8, e1002700.

Ekundayo, F.  O., Olukunle, O.F., and Ekundayo, E. A.(2012).  Biodegradation of Bonnylight crude oil bylocally isolated fungi from oil contaminated soils in Akure, Ondo state.  Malaysian Journal of Microbiology.

Linder, M.B., Szilvay, G.R., Nakari-Setälä, T., and Penttilä,M.E. (2005).  Hydrophobins: the proteinamphiphiles of filamentous fungi.  FEMS MicrobiolRev 29, 877–896.

Luke, A.K., and Burton, S.G. (2001).  A novel application for Neurospora crassa: Progress from batch culture to a membrane bioreactor for the bioremediation of phenols.  Enzyme and Microbial Technology 29,348–356.

Journal of Biological Sciences at Rutgers Camden (JBS) is licensed under a Creative Commons Attribution-NonCommercial-ShareAlike 4.0 International License