Open access peer-reviewed chapter

The Oleaginous Red Yeast Rhodotorula/Rhodosporidium: A Factory for Industrial Bioproducts

Written By

Mathew Lyman, Salustra Urbin, Cheryl Strout and Bonnee Rubinfeld

Submitted: 26 October 2018 Reviewed: 03 January 2019 Published: 05 February 2019

DOI: 10.5772/intechopen.84129

From the Edited Volume

Yeasts in Biotechnology

Edited by Thalita Peixoto Basso

Chapter metrics overview

1,829 Chapter Downloads

View Full Metrics


Rhodotorula genus, amended in 2015, is polyphyletic and contains Rhodotorula species that grow as single-cell yeast (monomorphic) and reproduce asexually via budding/fission (anamorphic); it also contains Rhodosporidium species that reproduce sexually (teleomorphic) and alternate between a yeast phase and dikaryotic filamentous phase (dimorphic). Several species of these “red yeast” produce industrial bioproducts, namely biofuel feedstocks, carotenoids, enzymes, and biosurfactants. This chapter highlights the biotechnology areas that Rhodotorula/Rhodosporidium contributes to and the future market value of those industries. The primary yeast species to be discussed include Rhodosporidium toruloides, Rhodotorula glutinis, Rhodosporidium diobovatum, Rhodosporidium kratochvilovae, Rhodotorula graminis, Rhodotorula babjevae, and Rhodotorula taiwanensis.


  • Rhodotorula
  • Rhodosporidium
  • red yeast
  • oleaginous
  • carotenoids
  • biofuels
  • biodiesel
  • antioxidants
  • DAAO enzyme
  • PAL enzyme
  • PEFA
  • surfactants
  • antagonistic yeast

1. Introduction

The aim of this chapter is to discuss the Rhodotorula genus in the context of biotechnology; it is not meant to be an academic “deep-dive” into all things known about the topic (their discovery and classification in the early 1900s could be a separate chapter unto itself [1]). The word “oleaginous” used in the chapter title is important, and it means “oil producing” or “rich in oils.” Much of the interest in these yeast stems from the fact that they often store excess carbon as triacylglycerols (TAG), not as polysaccharides [2]. These lipids can account for up to 70% of the cells dry mass depending on growth conditions, and these oils can be harvested and used as raw material in second-generation biodiesel production. Some Rhodotorula species also produce glycolipids, known as polyol esters of fatty acids (PEFA), which have broad interest in the biosurfactant industry. Several emerging publications and patents have been reported in this area. In addition, these yeasts are indeed “pinkish red.” These hues are carotenoid compounds produced by the yeast; these natural dyes can be extracted from Rhodotorula/Rhodosporidium and used in the food and vitamin industries. This chapter will further expand upon these intriguing facts and illustrate other examples that are in the scientific and patent literature, all while highlighting the future market potential of these “industrious” yeast.

1.1 A brief introduction to the Rhodotorula genus

There was a major revision of the subphylum Pucciniomycotina (Phylum Basidiomycota, Kingdom Fungi) published by Wang et al. in 2015; therefore, this chapter will focus solely on species that are categorized under the revised Rhodotorula genus in the Sporidiobolaceae family. Species to be discussed include Rhodosporidium toruloides, Rhodotorula glutinis, Rhodosporidium diobovatum, Rhodosporidium kratochvilovae, Rhodotorula graminis, Rhodotorula babjevae, and Rhodotorula taiwanensis. Species that were removed from the Rhodotorula genus in 2015, for example, Rhodotorula bogoriensis (reclassified as Pseudohyphozyma bogoriensis), will not be discussed in detail, even though it produces sophorolipid biosurfactants and is of industrial interest [3, 4, 5].

It is noteworthy that the Rhodotorula genus is polyphyletic. It contains Rhodotorula species that grow as single-cell yeast (monomorphic) and reproduce asexually via budding/fission (anamorphic) [6]. It also contains Rhodosporidium species that reproduce sexually (teleomorphic) and alternate between a yeast phase and dikaryotic filamentous phase (dimorphic). For Rhodosporidium, sexual reproduction begins with the fusing of compatible haploid yeast cells. They then grow as dikaryotic hyphae/mycelium, during which diploid teliospores are produced. Teliospores germinate forming basidium (where karyogamy and meiosis occur) and then extrude haploid basidiospores. Germination of basidiospores then restores the yeast phase of growth [7, 8]. Thus, the yeast phases of Rhodotorula and Rhodosporidium species are virtually indistinguishable. It is notable that “all known Rhodosporidium species [have been] isolated as haploid yeasts and have a bipolar mating behavior, i.e., their strains belong to either one of two complementary mating types, designated A1 and A2 or A and a” [9]. Due to this mating behavior, Rhodosporidium toruloides is being developed as an alternative biotechnology platform to Saccharomyces cerevisiae [10] with unique biochemical pathways for the production of biofuels, carotenoids, and industrial enzymes.

1.2 The industrial markets of the genus Rhodotorula

The most relevant question for the purposes of this chapter, regardless of the details of any given species, is “what biotechnology markets are impacted by the Rhodotorula genus?” Figure 1 summarizes the five major industrial markets where Rhodotorula yeast is utilized (or will likely be utilized in the future). These include biofuels, carotenoids, enzyme production (e.g. D-Amino acid oxidase (DAAO) and L-Phenylalanine ammonia lyase (PAL)), biosurfactants, and antagonistic yeast.

Figure 1.

The primary biotechnology industries impacted by the Rhodotorula genus. “Rhodotorula in Industry” was written in Rhodotorula taiwanensis MD1149 on a Hommel minimal salt (HMS) agar plate supplemented with glucose.

1.2.1 Biofuels

Biofuels continue to grow as a global industry, despite the challenges in production capacity and automotive engine compatibility [11]. The biofuels market size is expected to reach USD ~218 billion by 2022 [12]. For example, United Airlines announced in 2018 that it would begin blending more biofuel with conventional fuel, with a goal to reduce greenhouse emissions by 50% on all flights by 2050. It is expected that other airlines will follow suit, further increasing biofuel demand. Thus, there continues to be a need for microbial factories to produce biofuels [13, 14], especially given the market competition of using vegetable oils in both the food and biodiesel sectors [15]. Microorganisms may contribute to biofuel production in several ways; bisabolene (a diesel alternative) can be produced through bioengineering the isoprenoid pathway [16], and alkanes and alkenes can be produced through fatty acid biosynthesis inside the cell [15, 17]. Furthermore, microbial triglycerides, fatty acids, and other lipids can be converted to fatty acid methyl esters (FAME) by transesterification with methanol; FAME molecules are the primary component in biodiesel fuel.

The biofuels industry is largely divided between two types of fuel: biodiesel and bioethanol. These biofuels are further separated as “first generation, second generation, third generation, or fourth generation.” First-generation biofuels are produced using feedstocks that can also be used for food (e.g. soy, canola, or sunflower oils). These oils are transesterified into FAME by reacting a triacylglyceride with a short alcohol—usually methanol—in the presence of a catalyst (strong base) and heat. By contrast, second-generation biofuels are produced using non-food/non-edible sources such as wood, organic waste, and food crop waste. Second-generation biofuels are preferred as they do not utilize food crops for fuel. Third-generation biofuels are produced by engineered algae; fourth-generation biofuels are aimed at using biomass that captures CO2 while producing lipids that can be converted into biofuel. In this context, Rhodotorula/Rhodosporidium species that are used in biodiesel production are classified as “second generation” biodiesel producers that accumulate single-cell oils (SCOs); they will be discussed later in this chapter.

1.2.2 Carotenoids

In addition to lipids, these yeasts also produce carotenoids, which are of industrial interest. A recent report entitled “Global Carotenoid Market – Growth, Trends, and Forecast (2018–2023)” predicted a global carotenoid market of USD 2 billion by 2023 [18]. Carotenoids are yellow, orange, and red pigments produced by microorganisms, algae, higher plants, and some animals; these compounds are responsible for giving Rhodotorula species their “Red Yeast” moniker. Common examples of carotenoids include alpha-carotene, beta-carotene, beta-cryptoxanthin, lutein, lycopene, torulene, and torularhodin. Carotenoids are effective “quenchers” of reactive oxygen species (ROS), and the protective health effects of dietary carotenoids (as antioxidants) are of intense interest to the industrial research community [19]. Some carotenoids found in fruits and vegetables can be converted into vitamin A, namely beta-carotene, and to a lesser extent alpha-carotene and beta-cryptoxanthin (note: these compounds are often referred to as “provitamin A”). The impact of Vitamin A on human health is nicely summarized by Ulbricht et al. [20]. Carotenoids may also be used as natural coloring agents in the food, cosmetic, and pharmaceutical industries [21].

1.2.3 Enzymes

The Rhodotorula/Rhodosporidium enzymes, D-Amino acid oxidase (DAAO) and L-Phenylalanine ammonia lyase (PAL), have notable utility within industry. DAAO is a “FAD-dependent oxidoreductase that catalyzes stereospecifically the oxidative deamination of d-amino acids to α-keto acids, ammonia, and hydrogen peroxide” [22]. Even though DAAOs have been identified in a wide range of living organisms (e.g. bacteria, fungi, humans) [23], DAAO from certain Rhodotorula strains—discussed later in this chapter—has advantages over others, mainly due to a higher turnover rate and their increased stability of FAD binding [24]. This makes them ideal for the enzymatic deamination of cephalosporin C to 7-(5-oxoadipoamido)-cephalosporanic acid, an important intermediate for the production of cephalosporin [25]. Cephalosporin antibiotics are the largest selling class of antibiotics in a market, which is expected to reach USD 57 billion by 2024 [26, 27]. The PAL enzyme is also a major contributor to the enzymatic synthesis of “industrially relevant biomolecules,” namely pure L-phenylalanine (L-Phe), L-phenylalanine methyl ester (L-PM), and para-hydroxycinnamic acid (p-HCA) [28]. L-Phe, being an essential amino acid, is used in food formulations, feed for livestock, dietary supplements, and nutraceuticals [29]. L-PM is a precursor in the production of the sweetener aspartame [28], and p-HCA has utility in the cosmetic, health, and flavoring industries [30].

1.2.4 Biosurfactants

The biosurfactant market is estimated to be worth over USD 2.7 billion by 2024, primarily driven by their usage in the personal care and cosmetic industries [31]. Biosurfactants are small molecules that contain both hydrophilic and hydrophobic moieties and reduce the surface tension between oil and water mixtures (colloquially they are known as “green detergents” or “bio-soaps”); they are produced by several microbial species, viz., bacteria, yeast, and fungi [32, 33], and they are utilized extensively in multiple industries: cosmetics, food, explosive, pharmaceutical, detergents, and paints [34, 35]. The primary market barrier for biosurfactants is the vast library of existing chemical surfactants produced from petroleum feedstocks; end users can choose from hundreds of synthetic surfactants to fulfill their industry need. A major drawback is that chemical/synthetic surfactants can accumulate in the environment and are toxic to microbes, plants, aquatic life, and higher vertebrates including humans [36]. Therefore, because biosurfactants are biodegradable and petroleum-independent, their value will likely increase as pollution increases.

1.2.5 Antagonistic yeast

Rhodotorula species also play a role in the market of biocontrol agents, also known as antagonistic yeast. Significant losses in harvested fruit occur from decay by filamentous fungi such as Botrytis cinerea and Penicillium expansum [37]. In order to replace or augment chemical fungicides, biological control agents have been studied heavily the past two decades, with some yeast-based biocontrol products commercially available for specific commodities [38]. The ability to control different rots, on different fruits/vegetables, is still an important goal when developing postharvest biocontrol products. Therefore, antagonistic yeasts continue to be an active area of research, and several Rhodotorula species have shown promise in this field.


2. Species utilized in biotechnology

2.1 Rhodosporidium toruloides

Rhodosporidium toruloides has tremendous potential as a workhorse for multiple industrial applications [39]. It can grow on a variety of carbon sources associated with modern waste streams, and its bioproducts can be used in antibiotic manufacturing, biofuel synthesis, and the food industry.

R. toruloides is a nonpathogenic, aerobic, oleaginous red yeast that has been isolated from a variety of sources, e.g. conifers, soil, wood pulp, dry leaves, and a salt farm [2]. It is able to accumulate lipids to more than 70% of its dry cell weight [40, 41, 42, 43]; this occurs when carbon is in excess during growth, and other key nutrients such as nitrogen are sparse [44]. This lipid production can occur on a variety of different carbon sources, e.g. sugarcane juice, crude glycerol, lignocellulosic hydrolysates, vegetable market waste, and Jerusalem artichoke plants [45, 46, 47, 48, 49].

Due to the ability of R. toruloides to use different types of carbon sources for growth and lipid production, several studies have been carried out on growing this organism using “cheap” carbon sources that are the waste streams from other large industries, namely food waste and crude glycerol waste. Food wastes are an appealing nutrient source, because they are cheap, abundant, and decrease the environmental impact of disposing of these materials using traditional consumer and agricultural waste infrastructure [50, 51]. Crude glycerol is of interest as a “waste” source as the biofuel industry generates ~10% w/w of glycerol for every batch of biodiesel produced; R. toruloides has been shown to use crude glycerol as a carbon source for the production of microbial lipids of interest, even in the presence of impurities [52, 53]. In addition, it has been reported that R. toruloides has the ability to degrade/utilize hydrocarbon fuels [54]. This opens the possibility of using this yeast as a bioremediation tool for oil contaminated soil.

The bioproducts produced by R. toruloides are also of intense interest, namely lipids, enzymes, and carotenoids. Wild-type and engineered strains of R. toruloides are truly oleaginous. They produce lipids at high titer making them promising organisms for the production of lipid-based chemicals such as biofuels, lubricants, surfactants, solvents, waxes, creams, and adhesives [2, 55, 56]. It has been shown that under low nitrogen conditions, growth on simple sugars, like glucose, fructose, xylose, or the carbohydrate glycerol, can increase TAG production [55]. To this end, several groups are working to further increase lipid production using engineered strains of R. toruloides, primarily for biofuels [45, 56, 57]. As a practical example, scientists in Brazil have performed a successful diesel engine test using biodiesel manufactured from lipids produced by R. toruloides when the organism was grown in sugarcane juice (carbon source) and urea (nitrogen source) [48].

R. toruloides is also a “gold mine” for industrial enzymes. It produces a high titer of esterase enzymes, which makes this organism of immense interest to the drug industry. For example, the process of making antibiotics in the cephalosporin class of compounds requires 3-acyloxymethyl cephalosporins to be enzymatically deacylated to the more stable intermediary 3-acyloxymethyl cephalosporins. This can be achieved by the esterase enzymes produced by R. toruloides. Another enzyme produced by R. toruloides is phenylalanine ammonia-lyase (PAL), which is a major contributor to the enzymatic synthesis of pure L-phenylalanine (L-Phe), L-phenylalanine methyl ester (L-PM), and para-hydroxycinnamic acid (p-HCA) [28].

R. toruloides has also played an important role in the development of an enzyme substitution treatment of Phenylketonuria (PKU) [58, 59], a genetic mutation manifesting in the inability to metabolize L-Phe. In the human diet, L-phenylalanine is found naturally in protein foods, such as eggs, meats, fish, cheese, and soybeans, and is also produced as a product that can be added to foods, in the case of aspartame. The buildup of L-Phe in the blood is extremely toxic and impacts neurological function in the form of seizures, tremors, and loss of muscle coordination, especially in the extremities [58]. Newborns are screened for PKU at birth to increase the chances of early discovery and attempt to limit the effects of PKU. A new drug, Palynziq™, was developed based on some of the research that involved R. toruloides along with other PAL producing species; it was approved by the FDA in May 2018 as an enzyme substitution therapy for PKU [59].

2.2 Rhodotorula glutinis

Rhodotorula glutinis, like R. toruloides, is of high industrial importance as it also synthesizes numerous valuable compounds: lipids (SCO, single-cell oils), enzymes (in particular, PAL), and carotenoids (lycopene, β-carotene, torulene, and torularhodin). An example workflow of harvesting these bioproducts from these species is shown in Figure 2.

Figure 2.

An example workflow of harvesting bioproducts from members of the Rhodotorula genus.

R. glutinis isolates have been isolated from air, fruit, seawater, soil, grass, milk, and cheese products [60, 61]. Previously, 370 collection strains were assigned to this species based upon color and growth properties; however, additional methods—namely genetic sequencing—have pared this list to a few dozen strains [61]. The majority of these strains are spherical, ellipsoidal, or elongated in shape; aerobic; and mesophilic, although some can thrive under lower temperatures. They use different sources of carbon for growth: glucose, galactose, sucrose, maltose, trehalose, ethanol, glycerol, and hexadecane. R. glutinis lack the capacity to perform sugar fermentation and can grow in salt (10% NaCl) but cannot tolerate high sugar (>50% glucose). Depending on strain and growth conditions, their colonies are smooth, moist, and mucoid. As with most species in the Rhodotorula genus, color can be creamy, yellow, salmon, pink, orange coral, and blood red due to the production of carotenoids [60].

R. glutinis strains also produce several microbial oils—dominated by oleic, linoleic, palmitic, and stearic acid [62, 63], and the lipid content in their biomass can reach up to 72% [64]. Numerous factors can affect the lipid content and distribution including choice of strain, carbon sources (molasses, glucose, sucrose, glycerol, or waste materials), nitrogen sources (ammonium sulfate or chloride, yeast extract, or monosodium glutamate wastewater), C/N ratio, and cultivation times. These factors result in a wide lipid content distribution between 18 and 66% [60]. Additionally, investigators have worked to increase microbial oil production by either improving culture medium (Chinese patent CN102559788A) or by genetically engineering R. glutinis (Chinese patent CN102796675B).

It is noteworthy that several conditions were evaluated for the co-production of both lipid and carotenoids; variables tested included irradiation (as a stressor), temperatures, and C/N ratios. This allowed for the development of a two-stage cultivation strategy where the first stage maximized biomass and carotenoid production under irradiation/high temperature and then maximized lipid content when switched to dark/low temperature [63]. Additional growth studies have evaluated the effect of pH with potato wastewater and glycerol [65], modulating air flow rates using glycerol with yeast extract as the nutrient supply without pH control [66] and using lignocellulosic biomass [67]. Other parameters for scale-up were explored using an airlift bioreactor with mixed carbon sources [68]. R. glutinis has also been genetically engineered by introducing both the beta-carotene biosynthesis genes and cellulase genes to increase yields and co-production [69].

Per other industrial applications, R. glutinis has been evaluated as a biocontrol agent for post-harvest microbial diseases of fruit (US patent US5525132A). To this end, it has been shown to significantly reduce the incidence of the gray mold, Botrytis cinerea on strawberries and apples [70, 71] possibly due to the attachment capability of the antagonistic R. glutinis to B. cinerea [72]. R. glutinis was also used in combination with rhamnolipids to be more efficacious against Alternaria alternata infection in cherry tomato fruit than either agent alone [73].

It is also interesting that R. glutinis has been isolated during the production of olive oil and during the fermentation and storage of green and black olives in the USA [74, 75]. Olive processing results in the generation of large quantities of olive mill wastewater (OMW) high in phenol. Advantageously, R. glutinis has been utilized to treat OMW by dephenolization [76].

2.3 Rhodosporidium diobovatum

Rhodosporidium diobovatum has been identified as being a top-tier lipid producer among a group of 69 varied oleaginous yeast strains when glucose is used as the sole carbon source; it is also amenable to scalability [77, 78]. However, biofuel research has also focused on R. diobovatum as a second-generation biodiesel producer due to its ability to consume the impure glycerol waste that is created as a byproduct of first-generation biodiesel production [77]. For example, it was recently reported that R. diobovatum could be an “effective strain for production of neutral lipids” given its high yields of oleic, palmitic, and linoleic acid [79], although growth on glucose produced more TAGs than glycerol. Importantly, a glycerol consumption strategy would allow for the continued use and optimization of the first-generation biodiesel production pipelines, while simultaneously allowing investment in the development of second-generation approaches.

Interestingly, it has been reported that up to 70% of the cost to make biodiesel could be accrued after the production of the cell biomass, when chemical methods of extracting TAGs are used [80]. This is a costly process that requires the complete drying of biomass, full chemical diffusion of solvent into the cells [81], yeast cell wall disruption, and lipid extraction [2]. The direct conversion of wet biomass to biodiesel is the ideal solution, but current methods for that are not scalable, require high temperatures, have lengthy reaction periods, or utilize expensive catalysts [82]. Recent advances with microalgae utilized an ionic liquid for wet cell disruption and lipid extraction in less than 1.5 hours [83]. This same method was effective for R. diobovatum and was optimized to produce 97.1% conversion of maximum FAME yields in 2.5 hours at 65°C. They did note a loss of ~40% of their KOH catalyst but conclude that switching from a homogenous to a heterogeneous catalyst could mitigate this [80]. Therefore, research is ongoing to develop R. diobovatum as top biofuel production species.

R. diobovatum is also being examined in several novel applications. These include as a way to produce the vitamin supplement glutathione (GSH) [84], which has reported antioxidant properties, and also can be utilized as an anti-toxicant, as a cell metabolism modulator, and potentially as a neuromediator [85]. GSH is synthesized by canonical yeasts but was found in R. diobovatum using high performance liquid chromatography (HPLC). Consequently, the R. diobovatum synthase genes have been identified and characterized, with the expectation that it could be a valuable industrial producer of GSH in future [84]. This species also has potential as a bioremediation agent for fertilizer pollution. It efficiently assimilates nitrogen at higher rates than other yeasts and may become a useful tool for treating agriculture wastewater [86].

2.4 Rhodosporidium kratochvilovae and Rhodotorula graminis

Rhodosporidium kratochvilovae cultures have been used to create single-cell oil biodiesel when cultured on cane molasses, a sugar refinery waste product [87]. This biodiesel meets standard specifications in Europe and the USA for quality and purity (ASTM D6751, EN14214). The strain was originally discovered in a screen for oleaginous yeasts in Ethiopia. Of the 340 yeast isolates screened, 18 tested positive for oil production, and a R. kratochvilovae strain was one of three chosen as best candidates for further optimization based on productivity [88]. Multiple parameters of its cultivation were optimized for lipid production [89] prior to being used to create SCO.

Rhodotorula graminis has also been developed for biodiesel using crude glycerol and undetoxified lignocellulosic hydrolysate (hydrolyzed corn stover; aka, the stalks, leaves, and cobs that remain in fields after harvest) [90]. This strain produced a high lipid content (34% w/w) on hydrolysate and increased this production on waste crude glycerol (54% w/w) [90]. Further optimization of this production pipeline was discussed in patent US9322038B2 granted in 2016 to Washington State University for “Simultaneous saccharification and fermentation (SSF) of lignocellulosic biomass for single-cell oil production by oleaginous microorganisms,” with details of strains and culture conditions therein.

R. kratochvilovae and R. graminis also produce carotenoids (specifically beta-carotene, torularhodin, and torulene) in order to mitigate cellular damage done by ROS [91]. The benefits of beta-carotene have been understood for some time (antioxidant), but those of torularhodin and torulene only been elucidated recently, namely their potent anti-cancer [92] and anti-microbial activity [93]. Importantly, R. graminis and other Rhodotorula species have the ability to produce these rare carotenoids at high levels [94] and represent viable options for industrial production of these compounds.

R. kratochvilovae and R. graminis are also involved in the biological control of plant pathogens [95]. In a screen for yeasts that control pathogenic fungal strains, R. kratochvilovae LS11 was found to have high antagonistic activity [96]. Further development of this strain is hoped to be achieved, and its detoxification pathways are currently being examined in detail [97]. R. graminis was also been examined for this application, alone and in combination with fungicide [98].

The PAL enzyme of R. graminis (RgrPAL) has industrial interest as it is highly stable in comparison to that of other species, and the organism has flexible culturing requirements. Mutagenesis was employed to create mutants that expressed higher levels of PAL, including an isolate that showed a fourfold increase in production [99]. RgrPAL is also useful to pharmaceutical production, as it accepts analogues of its substrate L-Phe, which can be included in peptidomimetic drugs. The enzymatic activity of RgrPAL was recently increased 28-fold using directed evolution methodology in a recent study [100].

2.5 Rhodotorula babjevae and Rhodotorula taiwanensis

It was recently published that strains of Rhodotorula babjevae, and other Rhodotorula species, produce polyol esters of fatty acids (PEFA) [101, 102], similar in composition to extracellular glycolipids reported in the 1960s [103]. The authors stated that “discovery of these PEFA-secreting yeasts may aid in improving production of renewable, sustainable, environmentally friendly surfactants for use in household and industrial cleaning products, as well as many other applications” [102]. To this end, a full patent application was submitted in 2017 by the University of California for “Methods of producing polyol lipids,” with details of Rhodotorula strains and culture conditions that optimize PEFA production (WO2017184884A1).

It is important to note that the general utility of surfactants in industry—whether they be chemical surfactants or biosurfactants—is determined by their hydrophilic-lipophilic balance (HLB). The HLB concept was first published in 1948 brochure by Atlas Powder Company, along with a follow-up journal paper by William Griffin (an Atlas chemist) [104]. Griffin described HLB as follows: “emulsifiers consist of a molecule that combines both hydrophilic and lipophilic groups and the balance of the size and strength of these two opposite groups is called HLB. For the purpose of convenience, the effective balance of these groups is assigned a numeric value [105].” Griffin then went on to develop the HLB number system, on a scale 0–20, based on polyoxyethylene (POE)-type surfactants. Although the correct classification of surfactants in the HLB system is still being refined and debated [104, 106, 107], the concept of different surfactants having different HLB values remains the benchmark for surfactant science. An example of different surfactant HLB values, their solubility, applications, and use in industry is provided in Table 1.

Surfactant HLBSolubilityApplicationsIndustry examples
1.5–3Oil solubleAntifoaming agents (defoamers)Hydraulics, paper, oil drilling, machine tools
3–6Oil solubleWater-in-oil emulsionsCosmetics, sunscreen, margarine
7–9Oil solubleWetting and spreading agentsHerbicides, fertilizers
12–16Water solubleOil-in-water-emulsionsMayonnaise, cosmetics, dispersants
13–15Water solubleDetergentsLaundry and dishwashing detergents
15–18Water solubleSolubilizing agentsPharmaceuticals

Table 1.

The hydrophilic-lipophilic balance (HLB) scale covers 0–20, with different industrial applications depending on the HLB value.

Perhaps the greatest market barrier that biosurfactants face is their limited coverage of the HLB scale. Therefore, they can only be used in very targeted commercial applications. By contrast, hundreds of petroleum-derived chemical surfactants conveniently cover the entire HLB scale, to such an extent that their massive usage is now poisoning the environment [36]. Biosurfactants offer a more environmentally friendly option (biodegradable and petroleum-independent); however, there are a limited number of commercially available biosurfactants, and each one covers only a small portion of the HLB scale.

Therefore, in order to expand PEFA coverage on the HLB scale, it was recently published that a new strain of Rhodotorula taiwanensis produced hypoacetylated PEFA compounds compared to those produced by Rhodotorula babjevae [108, 109]; the difference in their acetylation profiles resulted in different surface tension of the growth medium, i.e. a different hydrophilic-lipophilic balance. It was noted in a subsequent patent filing that although the current Rhodotorula taiwanensis strain produced a complex mixture of acetylated and non-acetylated biosurfactants, it could be genetically engineered to produce an unmodified (non-acetylated) “base compound.” This base compound could then be “tuned” to produce the full range of biosurfactants that could more extensively cover the HLB scale (Figure 3).

Figure 3.

A “tunable” PEFA biosurfactant. Rhodotorula taiwanensis would be genetically engineered to produce a single PEFA “base compound” and then be systematically modified to move up and down the HLB scale.

This “tunable” biosurfactant approach, using a single species of PEFA compound that can be systematically modified, provides a viable option for competing directly with all types of chemical surfactants, across all market sectors. The biosurfactant’s chemical topology, highlighted by the carbohydrate unit’s hydroxyl groups, offers further opportunity for chemical modifications that have a direct impact on the physical properties of the surfactant (e.g. antifoaming, emulsifying, wetting, detergent, and solubilizing). Each hydroxyl (OH) group on the molecule has the ability to be acetylated, sulfated, or phosphorylated, leading to modified versions of the parent molecule with different physicochemical properties. Therefore, modification of the hydroxyl groups by chemical and/or biochemical means will enable scientists to produce various biosurfactant products best suited for their unique industrial applications—serving markets not currently served by existing biosurfactants.


3. Conclusions

The Rhodotorula genus currently plays a significant role in yeast biotechnology and is poised to expand into various industrial markets: biofuels, carotenoids, biocontrol agents, enzymes, bioremediation, cosmetics, and others. Several advanced genetic systems are currently being developed for these red yeast, and they will likely become an alternative biotechnology platform to Saccharomyces cerevisiae [10]. The future looks bright (red) for these fascinating, diverse, and versatile oleaginous microbes.



This work was performed under the auspices of the U.S. Department of Energy by Lawrence Livermore National Laboratory under Contract DE-AC52-07NA27344. We thank Kristin Lyman for assistance with the graphic design.


Conflict of interest

The authors declare no conflict of interest.


  1. 1. Hasegawa T, Banno I, Yamauchi S. A taxonomic study on the genus Rhodotorula. The Journal of General and Applied Microbiology. 1960;6(3):196-215
  2. 2. Ageitos JM, Vallejo JA, Veiga-Crespo P, Villa TG. Oily yeasts as oleaginous cell factories. Applied Microbiology and Biotechnology. 2011;90(4):1219-1227
  3. 3. Nunez A, Ashby R, Foglia TA, Solaiman DK. LC/MS analysis and lipase modification of the sophorolipids produced by Rhodotorula bogoriensis. Biotechnology Letters. 2004;26(13):1087-1093
  4. 4. Ribeiro IA, Bronze MR, Castro MF, Ribeiro MH. Design of selective production of sophorolipids by Rhodotorula bogoriensis through nutritional requirements. Journal of Molecular Recognition: JMR. 2012;25(11):630-640
  5. 5. Solaiman DK, Ashby RD, Crocker NV. High-titer production and strong antimicrobial activity of sophorolipids from Rhodotorula bogoriensis. Biotechnology Progress. 2015;31(4):867-874
  6. 6. Morrow CA, Fraser JA. Sexual reproduction and dimorphism in the pathogenic basidiomycetes. FEMS Yeast Research. 2009;9(2):161-177
  7. 7. Boekhout T, Fonseca Á, Sampaio JP, Bandoni RJ, Fell JW, Kwon-Chung KJ. Discussion of teleomorphic and anamorphic basidiomycetous yeasts. In: The Yeasts: A Taxonomic Study. London, UK: Elsevier; 2011
  8. 8. Fell JW, Boekhout T, Fonseca A, Sampaio JP. Basidiomycetous yeasts. In: DJ ML, EJ ML, Lemke P, editors. The Mycota VII Part B. Systematics and Evolution. Berlin, Heidelberg: Springer-Verlag; 2001
  9. 9. Coelho MA, Rosa A, Rodrigues N, Fonseca A, Goncalves P. Identification of mating type genes in the bipolar basidiomycetous yeast Rhodosporidium toruloides: First insight into the MAT locus structure of the Sporidiobolales. Eukaryotic Cell. 2008;7(6):1053-1061
  10. 10. Kumar S, Kushwaha H, Bachhawat AK, Raghava GP, Ganesan K. Genome sequence of the oleaginous red yeast Rhodosporidium toruloides MTCC 457. Eukaryotic Cell. 2012;11(8):1083-1084
  11. 11. Hassan MK, A. An overview of biofuel as a renewable energy source: Development and challenges. Procedia Engineering. 2013;56:39-53
  12. 12. Research ZM. Biofuels market analysis by type (bioethanol, biodiesel), and by form (solid, liquid, and gaseous)—Global industry perspective, comprehensive analysis, and forecast, 2016-2022. 2018.
  13. 13. Lee SY, Kim HM, Cheon S. Metabolic engineering for the production of hydrocarbon fuels. Current Opinion in Biotechnology. 2015;33:15-22
  14. 14. Peralta-Yahya PP, Zhang F, del Cardayre SB, Keasling JD. Microbial engineering for the production of advanced biofuels. Nature. 2012;488(7411):320-328
  15. 15. Zhou Y, Eduard K, Nielsen J. Barriers and opportunities in bio-based production of hydrocarbons. Nature Energy. 2018;3:925-935
  16. 16. Peralta-Yahya PP, Ouellet M, Chan R, Mukhopadhyay A, Keasling JD, Lee TS. Identification and microbial production of a terpene-based advanced biofuel. Nature Communications. 2011;2:483
  17. 17. Pfleger BF, Gossing M, Nielsen J. Metabolic engineering strategies for microbial synthesis of oleochemicals. Metabolic Engineering. 2015;29:1-11
  18. 18. Intelligence Mordor. Global Carotenoid Market—Growth, Trends, and Forecast (2018-2023). 2018
  19. 19. Fiedor J, Burda K. Potential role of carotenoids as antioxidants in human health and disease. Nutrients. 2014;6(2):466-488
  20. 20. Ulbricht C, Basch E, Chao W, Conquer J, Costa D, Culwell S, et al. An evidence-based systematic review of vitamin A by the natural standard research collaboration. Journal of Dietary Supplements. 2012;9(4):299-416
  21. 21. Mortensen A. Carotenoids and other pigments as natural colorants. Pure and Applied Chemistry. 2006;78(8):1477-1491
  22. 22. Liu Y, Koh CMJ, Ngoh ST, Ji L. Engineering an efficient and tight D-amino acid-inducible gene expression system in Rhodosporidium/Rhodotorula species. Microbial Cell Factories. 2015;14:170
  23. 23. Pollegioni L, Piubelli L, Sacchi S, Pilone MS, Molla G. Physiological functions of D-amino acid oxidases: From yeast to humans. Cellular and Molecular Life Sciences. 2007;64(11):1373-1394
  24. 24. Pilone MS. D-amino acid oxidase: New findings. Cellular and Molecular Life Sciences. 2000;57(12):1732-1747
  25. 25. Hsieh HC, Kuan IC, Lee SL, Tien GY, Wang YJ, Yu CY. Stabilization of D-amino acid oxidase from Rhodosporidium toruloides by immobilization onto magnetic nanoparticles. Biotechnology Letters. 2009;31(4):557-563
  26. 26. Hamad B. The antibiotics market. Nature Reviews. Drug Discovery. 2010;9(9):675-676
  27. 27. Research GV. Antibiotics Market Size To Reach $57.0 Billion By 2024. 2016
  28. 28. MacDonald MC, Arivalagan P, Barre DE, MacInnis JA, D'Cunha GB. Rhodotorula glutinis phenylalanine/tyrosine ammonia lyase enzyme catalyzed synthesis of the methyl ester of Para-hydroxycinnamic acid and its potential antibacterial activity. Frontiers in Microbiology. 2016;7:281
  29. 29. Ding D, Liu Y, Xu Y, Zheng P, Li H, Zhang D, et al. Improving the production of L-phenylalanine by identifying key enzymes through multi-enzyme reaction system in vitro. Scientific Reports. 2016;6:32208
  30. 30. Vargas-Tah A, Gosset G. Production of cinnamic and p-Hydroxycinnamic acids in engineered microbes. Frontiers in Bioengineering and Biotechnology. 2015;3:116
  31. 31. Global Market Insights I. Biosurfactants Market worth over $2.7 bn by 2024. 2018
  32. 32. Satpute SK, Banpurkar AG, Dhakephalkar PK, Banat IM, Chopade BA. Methods for investigating biosurfactants and bioemulsifiers: A review. Critical Reviews in Biotechnology. 2010;30(2):127-144
  33. 33. Uzoigwe C, Burgess JG, Ennis CJ, Rahman PK. Bioemulsifiers are not biosurfactants and require different screening approaches. Frontiers in Microbiology. 2015;6:245
  34. 34. Reis RS, Pacheco GJ, Pereira AG, Freire DMG. Biosurfactants: Production and applications. In: Rosenkranz RCF, editor. Biodegradation—Life of Science. Rijeka: InTech; 2013
  35. 35. Varvaresou A, Iakovou K. Biosurfactants in cosmetics and biopharmaceuticals. Letters in Applied Microbiology. 2015;61(3):214-223
  36. 36. Rebello S, Asok AK, Mundayoor S, Jisha MS. Surfactants: Chemistry, toxicity and remediation. In: Lichtfouse E et al., editors. Pollutant Diseases, Remediation and Recycling. Switzerland: Springer International Publishing; 2013
  37. 37. Castoria R, Filipo DC, Lima G, De Cicco V. β-1,3-glucanase activity of two saprophytic yeasts and possible mode of action as biocontrol agents against postharvest diseases. Postharvest Biology and Technology. 1997;12(3):293-300
  38. 38. Liu J, Sui Y, Wisniewski M, Droby S, Liu Y. Review: Utilization of antagonistic yeasts to manage postharvest fungal diseases of fruit. International Journal of Food Microbiology. 2013;167(2):153-160
  39. 39. Park YK, Nicaud JM, Ledesma-Amaro R. The engineering potential of Rhodosporidium toruloides as a workhorse for biotechnological applications. Trends in Biotechnology. 2018;36(3):304-317
  40. 40. Li Q , Du W, Liu D. Perspectives of microbial oils for biodiesel production. Applied Microbiology and Biotechnology. 2008;80(5):749-756
  41. 41. Singh G, Jawed A, Paul D, Bandyopadhyay KK, Kumari A, Haque S. Concomitant production of lipids and carotenoids in Rhodosporidium toruloides under osmotic stress using response surface methodology. Frontiers in Microbiology. 2016;7:1686
  42. 42. Zhu Z, Zhang S, Liu H, Shen H, Lin X, Yang F, et al. A multi-omic map of the lipid-producing yeast Rhodosporidium toruloides. Nature Communications. 2012;3:1112
  43. 43. Li Y, Zhao Z, Bai F. High-density cultivation of oleaginous yeast Rhodosporidium toruloides Y4 in fed-batch culture. Enzyme and Microbial Technology. 2007;41:312-317
  44. 44. Papanikolaou S, Aggelis G. Lipids of oleaginous yeasts. Part I: Biochemistry of single cell oil production. European Journal of Lipid Science and Technology. 2011;113:1031-1051
  45. 45. Fei Q , O'Brien M, Nelson R, Chen X, Lowell A, Dowe N. Enhanced lipid production by Rhodosporidium toruloides using different fed-batch feeding strategies with lignocellulosic hydrolysate as the sole carbon source. Biotechnology for Biofuels. 2016;9:130
  46. 46. Hu C, Zhao X, Zhao J, Wu S, Zhao ZK. Effects of biomass hydrolysis by-products on oleaginous yeast Rhodosporidium toruloides. Bioresource Technology. 2009;100(20):4843-4847
  47. 47. Singh G, Sinha S, Bandyopadhyay KK, Lawrence M, Paul D. Triauxic growth of an oleaginous red yeast Rhodosporidium toruloides on waste 'extract' for enhanced and concomitant lipid and beta-carotene production. Microbial Cell Factories. 2018;17(1):182
  48. 48. Soccol CR, Dalmas Neto CJ, Soccol VT, Sydney EB, da Costa ESF, Medeiros ABP, et al. Pilot scale biodiesel production from microbial oil of Rhodosporidium toruloides DEBB 5533 using sugarcane juice: Performance in diesel engine and preliminary economic study. Bioresource Technology. 2017;223:259-268
  49. 49. Zhao X, Wu S, Hu C, Wang Q , Hua Y, Zhao ZK. Lipid production from Jerusalem artichoke by Rhodosporidium toruloides Y4. Journal of Industrial Microbiology & Biotechnology. 2010;37(6):581-585
  50. 50. Pham TP, Kaushik R, Parshetti GK, Mahmood R, Balasubramanian R. Food waste-to-energy conversion technologies: Current status and future directions. Waste Management. 2015;38:399-408
  51. 51. Zeng Y, Xie T, Li P, Jian B, Li X, Xie Y, et al. Enhanced lipid production and nutrient utilization of food waste hydrolysate by mixed culture of oleaginous yeast Rhodosporidium toruloides and oleaginous microalgae Chlorella vulgaris. Renewable Energy. 2018;126:915-923
  52. 52. Uprety BK, Swaroop Dalli S, Rakshit S. Bioconversion of crude glycerol to microbial lipid using a robust oleaginous yeast Rhodosporidium toruloides ATCC 10788 capable of growing in the presence of impurities. Energy Conversion and Management. 2017;135:117-128
  53. 53. Xu J, Xuebing Z, Wang W, Du W, Liu D. Microbial conversion of biodiesel byproduct glycerol to triacylglycerols by oleaginous yeast Rhodosporidium toruloides and the individual effect of some impurities on lipid production. Biochemical Engineering Journal. 2012;65:30-36
  54. 54. Kumari M, Jayanthi A. Biodegradation of diesel oil using yeast Rhodosporidium toruloides. Research Journal of Environmental Toxicology. 2011;5(6):369-377
  55. 55. Zhang S, Ito M, Skerker JM, Arkin AP, Rao CV. Metabolic engineering of the oleaginous yeast Rhodosporidium toruloides IFO0880 for lipid overproduction during high-density fermentation. Applied Microbiology and Biotechnology. 2016;100(21):9393-9405
  56. 56. Zhang S, Skerker JM, Rutter CD, Maurer MJ, Arkin AP, Rao CV. Engineering Rhodosporidium toruloides for increased lipid production. Biotechnology and Bioengineering. 2016;113(5):1056-1066
  57. 57. Yaegashi J, Kirby J, Ito M, Sun J, Dutta T, Mirsiaghi M, et al. Rhodosporidium toruloides: A new platform organism for conversion of lignocellulose into terpene biofuels and bioproducts. Biotechnology for Biofuels. 2017;10:241
  58. 58. Gamez A, Sarkissian CN, Wang L, Kim W, Straub M, Patch MG, et al. Development of pegylated forms of recombinant Rhodosporidium toruloides phenylalanine ammonia-lyase for the treatment of classical phenylketonuria. Molecular Therapy. 2005;11(6):986-989
  59. 59. Levy HL, Sarkissian CN, Scriver CR. Phenylalanine ammonia lyase (PAL): From discovery to enzyme substitution therapy for phenylketonuria. Molecular Genetics and Metabolism. 2018;124(4):223-229
  60. 60. Kot AM, Blazejak S, Kurcz A, Gientka I, Kieliszek M. Rhodotorula glutinis-potential source of lipids, carotenoids, and enzymes for use in industries. Applied Microbiology and Biotechnology. 2016;100(14):6103-6117
  61. 61. Sampaio J. Rhodotorula Harrison. In: Kurtzman C, editor. The Yeasts: A Taxonomic Study. Amsterdam: Elsevier; 1928
  62. 62. Easterling ER, French WT, Hernandez R, Licha M. The effect of glycerol as a sole and secondary substrate on the growth and fatty acid composition of Rhodotorula glutinis. Bioresource Technology. 2009;100(1):356-361
  63. 63. Zhang Z, Zhang X, Tan T. Lipid and carotenoid production by Rhodotorula glutinis under irradiation/high-temperature and dark/low-temperature cultivation. Bioresource Technology. 2014;157:149-153
  64. 64. Meng XY, Jianming Y, Xu X, Zhang L, Nie Q , Xian M. Biodiesel production from oleaginous microorganisms. Renewable Energy. 2009;34:1-5
  65. 65. Kot AM, Błażejak S, Kurez A, Gientka I, Bzducha-Wrobel A, Maliszewska M, et al. Effect of initial pH of medium with potato wastewater and glycerol on protein, lipid and carotenoid biosynthesis by Rhodotorula glutinis. Electronic Journal of Biotechnology. 2017;27:25-31
  66. 66. Karamerou EE, Theodoropoulos C, Webb C. A biorefinery approach to microbial oil production from glycerol by Rhodotorula glutinis. Biomass and Bioenergy. 2016;89:113-122
  67. 67. Dai C-C, Jie T, Xie F, Dai Y-J, Zhao M. Biodiesel generation from oleaginous yeast Rhodotorula glutinis with xylose assimilating capacity. African Journal of Biotechnology. 2007;6(18):2130-2134
  68. 68. Yen H-W, Chang J-T, Chang J-S. The growth of oleaginous Rhodotorula glutinis in an internal-loop airlift bioreactor by using mixture substrates of rice straw hydrolysate and crude glycerol. Biomass and Bioenergy. 2015;80:38-43
  69. 69. Pi HW, Anandharaj M, Kao YY, Lin YJ, Chang JJ, Li WH. Engineering the oleaginous red yeast Rhodotorula glutinis for simultaneous beta-carotene and cellulase production. Scientific Reports. 2018;8(1):10850
  70. 70. Zhang H, Wang L, Dong Y, Jiang S, Cao J, Meng R. Postharvest biological control of gray mold decay of strawberry with Rhodotorula glutinis. Biological Control. 2007;40:287-292
  71. 71. Zhang H, Wang L, Ma L, Dong Y, Jiang S, Xu B, et al. Biocontrol of major postharvest pathogens on apple using Rhodotorula glutinis and its effects on postharvest quality parameters. Biological Control. 2009;48:79-83
  72. 72. Li B, Peng H, Tian S. Attachment capability of antagonistic yeast Rhodotorula glutinis to Botrytis cinerea contributes to biocontrol efficacy. Frontiers in Microbiology. 2016;7:601
  73. 73. Yan F, Xu S, Chen Y, Zheng X. Effect of rhamnolipids on Rhodotorula glutinis biocontrol of Alternaria alternate infection in cherry tomato fruit. Postharvest Biology and Technology. 2014;97:32-35
  74. 74. Arroyo-Lopez FN, Querol A, Bautista-Gallego J, Garrido-Fernandez A. Role of yeasts in table olive production. International Journal of Food Microbiology. 2008;128(2):189-196
  75. 75. Yousuf A, Sannino F, Addorisio V, Pirozzi D. Microbial conversion of olive oil mill wastewaters into lipids suitable for biodiesel production. Journal of Agricultural and Food Chemistry. 2010;58(15):8630-8635
  76. 76. Gaye B, Takaç S. Parameters and kinetics of olive mill wastewater dephenolization by immobilized Rhodotorula glutinis cells. Environmental Technology. 2014;35:3074-3081
  77. 77. Munch G. Characterization and Comparison of Different Oleaginous Yeasts and Scale-Up of Single-Cell Oil Production Using Rhodosporidium diobovatum. University of Manitoba; 2015
  78. 78. Sitepu IR, Sestric R, Ignatia L, Levin D, German JB, Gillies LA, et al. Manipulation of culture conditions alters lipid content and fatty acid profiles of a wide variety of known and new oleaginous yeast species. Bioresource Technology. 2013;144:360-369
  79. 79. Nasirian N, Mirzaie M, Cicek N, Levin DB. Lipid and carotenoid synthesis by Rhodosporidium diobovatum, grown on glucose versus glycerol, and its biodiesel properties. Canadian Journal of Microbiology. 2018;64(4):277-289
  80. 80. Ward V, Garret M, Cicek N, Rehmann L. Direct conversion of the oleaginous yeast Rhodosporidium diobovatum to biodiesel using the ionic liquid. ACS Sustainable Chemistry & Engineering. 2017;5:5562-5570
  81. 81. Halim R, Gladman B, Danquah MK, Webley P. Oil extraction from microalgae for biodiesel production. Bioresource Technology. 2011;102(1):178-185
  82. 82. Griffiths MJ, van Hille RP, Harrison ST. Selection of direct transesterification as the preferred method for assay of fatty acid content of microalgae. Lipids. 2010;45(11):1053-1060
  83. 83. VCA O, Plechkova NV, Seddon KR, Rehmann L. Disruption and wet extraction of the microalgae Chlorella vulgaris using room-temperature ionic liquids. ACS Sustainable Chemistry & Engineering. 2016;4:591-600
  84. 84. Kong M, Wang F, Tian L, Tang H, Zhang L. Functional identification of glutamate cysteine ligase and glutathione synthetase in the marine yeast Rhodosporidium diobovatum. Naturwissenschaften. 2017;105(1-2):4
  85. 85. Pompella A, Visvikis A, Paolicchi A, De Tata V, Casini AF. The changing faces of glutathione, a cellular protagonist. Biochemical Pharmacology. 2003;66(8):1499-1503
  86. 86. Civiero E, Pintus M, Ruggeri C, Tamburini E, Sollai F, Sanjust E, et al. Physiological and phylogenetic characterization of Rhodotorula diobovata DSBCA06, a Nitrophilous yeast. Biology (Basel). 2018;7(3):39
  87. 87. Jiru TM, Steyn L, Pohl C, Abate D. Production of single cell oil from cane molasses by Rhodotorula kratochvilovae (syn, Rhodosporidium kratochvilovae) SY89 as a biodiesel feedstock. Chemistry Central Journal. 2018;12(1):91
  88. 88. Jiru TM, Abate D, Kiggundu N, Pohl C, Groenewald M. Oleaginous yeasts from Ethiopia. AMB Express. 2016;6(1):78
  89. 89. Jiru TM, Groenewald M, Pohl C, Steyn L, Kiggundu N, Abate D. Optimization of cultivation conditions for biotechnological production of lipid by Rhodotorula kratochvilovae (syn, Rhodosporidium kratochvilovae) SY89 for biodiesel preparation. 3 Biotech. 2017;7(2):145
  90. 90. Galafassi S, Cucchetti D, Pizza F, Franzosi G, Bianchi D, Compagno C. Lipid production for second generation biodiesel by the oleaginous yeast Rhodotorula graminis. Bioresource Technology. 2012;111:398-403
  91. 91. Breierova E, Certik M, Marova I, Vadkertiova R. The effect of Zn(II) ions and reactive oxygen on the uptake of zinc and production of carotenoids by selected red yeasts. Chemistry & Biodiversity. 2018;15(6):e1800069
  92. 92. Du C, Li Y, Guo Y, Han M, Zhang W, Qian H. The suppression of torulene and torularhodin treatment on the growth of PC-3 xenograft prostate tumors. Biochemical and Biophysical Research Communications. 2016;469(4):1146-1152
  93. 93. Ungureanu C, Ferdes M. Evaluation of antioxidant and antimicrobial activities of Torularhodin. Advanced Science Letters. 2012;18:50-53
  94. 94. Kot AM, Blazejak S, Gientka I, Kieliszek M, Brys J. Torulene and torularhodin: "new" fungal carotenoids for industry? Microbial Cell Factories. 2018;17(1):49
  95. 95. Miccoli C, Palmieri D, De Curtis F, Lima G, Ianiri G, Castoria R. Complete genome sequence of the biocontrol agent yeast Rhodotorula kratochvilovae strain LS11. Genome Announcements. 2018;6(10):e00120-e00118
  96. 96. De Curtis F. I Lieviti Nella Lotta Biologica Contro Patogeni Fungini Degli Ortofrutticoli in Postraccolta: Attività e Meccanismi D'azione Coinvolti. Bologna, Italy: University of Bologna; 1998
  97. 97. Pinedo C, Wright SAI, Collado IG, Goss RJM, Castoria R, Hrelia P, et al. Isotopic labeling studies reveal the Patulin detoxification pathway by the biocontrol yeast Rhodotorula kratochvilovae LS11. Journal of Natural Products. 2018;81:2692-2699
  98. 98. Buck JW. Combinations of fungicides with Phylloplane yeasts for improved control of Botrytis cinerea on Geranium seedlings. Phytopathology. 2004;94(2):196-202
  99. 99. Orndorff SA, Costantino N, Stewart D, Durham DR. Strain improvement of Rhodotorula graminis for production of a novel l-phenylalanine Ammonia-Lyase. Applied and Environmental Microbiology. 1988;54(4):996-1002
  100. 100. Rowles I, Groenendaal B, Binay B, Malone KJ, Willies SC, Turner NJ. Engineering of phenylalanine ammonia lyase from Rhodotorula graminis for the enhanced synthesis of unnatural L-amino acids. Tetrahedron. 2016;72:7343-7347
  101. 101. Cajka T, Garay LA, Sitepu IR, Boundy-Mills KL, Fiehn O. Multiplatform mass spectrometry-based approach identifies extracellular glycolipids of the yeast Rhodotorula babjevae UCDFST 04-877. Journal of Natural Products. 2016;79(10):2580-2589
  102. 102. Garay LA, Sitepu IR, Cajka T, Fiehn O, Cathcart E, Fry RW, et al. Discovery of synthesis and secretion of polyol esters of fatty acids by four basidiomycetous yeast species in the order Sporidiobolales. Journal of Industrial Microbiology & Biotechnology. 2017;44:923-936
  103. 103. Tulloch A, Spencer J. Extracellular glycolipids of Rhodotorula species. Canadian Journal of Chemistry. 1964;42:830-835
  104. 104. Pasquali RC, Taurozzi MP, Bregni C. Some considerations about the hydrophilic-lipophilic balance system. International Journal of Pharmaceutics. 2008;356(1-2):44-51
  105. 105. Griffin W. Classification of surface-active agents by HLB. Journal of the Society of Cosmetic Chemists. 1949;1(5):311-326
  106. 106. Yamashita Y, Sakamoto K. Hydrophilic–lipophilic balance (HLB): Classical indexation and novel indexation of surfactant. In: Encyclopedia of Biocolloid and Biointerface Science. Hoboken, NJ, USA: John Wiley & Sons Inc; 2016
  107. 107. Davies J. London. 1957.
  108. 108. Lyman M, Rubinfeld B, Leif R, Mulcahy H, Dugan L, Souza B. Rhodotorula taiwanensis MD1149 produces hypoacetylated PEFA compounds with increased surface activity compared to Rhodotorula babjevae MD1169. PLoS One. 2018;13(1):e0190373
  109. 109. Tkavc R, Matrosova VY, Grichenko OE, Gostincar C, Volpe RP, Klimenkova P, et al. Prospects for fungal bioremediation of acidic radioactive waste sites: Characterization and genome sequence of Rhodotorula taiwanensis MD1149. Frontiers in Microbiology. 2017;8:2528

Written By

Mathew Lyman, Salustra Urbin, Cheryl Strout and Bonnee Rubinfeld

Submitted: 26 October 2018 Reviewed: 03 January 2019 Published: 05 February 2019