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Johnson Matthey Technol. Rev., 2021, 65, (2), 227

doi:10.1595/205651321x16024905831259

Catalytic Hydroprocessing of Single-Cell Oils to Hydrocarbon Fuels

Converting microbial lipids to fuels is a promising approach to replace fossil fuels

  • Jacob S. Kruger*
  • Renewable Resources and Enabling Sciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
  • Eric P. Knoshaug
  • Bioscience Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
  • Tao Dong
  • Renewable Resources and Enabling Sciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
  • Tobias C. Hull
  • Renewable Resources and Enabling Sciences Center, National Renewable Energy Laboratory, 15013 Denver West Parkway, Golden, CO 80401, USA
  • Philip T. Pienkos
  • Polaris Renewables, 906 South Cole Drive, Lakewood, CO 80228, USA
  • *Email: Jacob.Kruger@nrel.gov
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Article Synopsis

Microbial lipids hold great promise as biofuel precursors, and research efforts to convert such lipids to renewable diesel fuels have been increasing in recent years. In contrast to the numerous literature reviews on growing, characterising and extracting lipids from oleaginous microbes, and on converting vegetable oils to hydrocarbon fuels, this review aims to provide insight into aspects that are specific to hydroprocessing microbial lipids. While standard hydrotreating catalysts generally perform well with terrestrial oils, differences in lipid speciation and the presence of co-extracted compounds, such as chlorophyll and sterols, introduce additional complexities into the process for microbial lipids. Lipid cleanup steps can be introduced to produce suitable feedstocks for catalytic upgrading.

Introduction

Oils derived from oleaginous microbes, such as algae, yeast and bacteria (so-called single-cell oils (SCOs)) have long held promise as a biofuel precursor due to the high lipid content (>20%), high growth rate and the ability of these microbes to be cultivated on feedstocks or in areas that do not compete for terrestrial food production (1). Historically, the availability of microbial lipids has been quite limited, but technology developments in food and feed applications may begin to benefit lipid production for biofuels as well. Corbion produces an algae oil high in oleic acid for culinary use, and an algae-based omega-3 fatty-acid-enriched product for fish feed. Qualitas Health, DSM, Evonik and Veramaris also produce omega-3 enriched oils for food and feed applications, though to our knowledge only Qualitas Health is producing the algae photoautotrophically. While the target omega-3 fraction is not ideal for fuel production due to long carbon chains and numerous double bonds that would increase hydrogen consumption during hydroprocessing, the process to enrich omega-3 oils likely generates a byproduct of lighter and more saturated fatty acids that may be more suitable for fuel production than the omega-3 fraction. Additionally, although many oil majors and startup companies have pivoted away from algal biofuels, a partnership between Synthetic Genomics and ExxonMobil to improve algal biology for fuel production is still active. The increasing availability of cellulosic sugars may also pave the way for larger scale cultivation of heterotrophic oleaginous microbes.

While both the growth and characterisation of oleaginous microbes (113) and the conversion of vegetable oils to fuels (1327) have been extensively studied, there has been considerably less research on converting SCOs to fuels, and especially on converting SCOs to so-called ‘drop-in’ hydrocarbon fuels that are compatible with existing fuel infrastructure (2830). However, production of hydrocarbon fuels from SCOs is a growing area of research and hydroprocessing of these oils requires some considerations that are unique to the differing composition of SCOs. Enough recent studies have been published that a summary of findings and perspective on research directions is warranted, especially with respect to the differences between terrestrial oils and SCOs. In particular, lipid recovery, lipid cleanup and catalyst selection all appear to play a key role in hydroprocessing performance, and there are clear needs for deeper research in each of these areas.

Lipid Recovery

In most cases, SCOs are produced intracellularly, and thus necessitate separation from the other cell components prior to upgrading. Additionally, the microbial biomass is typically recovered with a very high water content (≥80%) relative to terrestrial biomass. Thus, while vegetable oils can be effectively extracted by pressing, solvent extraction or a combination thereof, these operations are usually not effective when applied directly to microbial biomass. Furthermore, it is energy intensive to dehydrate microbial biomass and therefore strategies that can extract oil in high yields from wet biomass are required. The first step is commonly to rupture the microbial cell wall which allows access to the lipids.

Various cell wall lysis methods have been applied for this application, such as high-pressure homogenisation (HPH), bead milling, ultrasound, pulsed electric field (PEF), osmotic shock, microwave, subcritical water hydrolysis, enzymatic hydrolysis, autolysis and chemical hydrolysis (31). From the perspective of energy consumption, pretreatment methods such as HPH, subcritical water hydrolysis, enzymatic and chemical hydrolysis are more attractive due to the relatively lower energy requirements. Moreover, all these methods can be scaled up to industrial application (32). Researchers at the National Renewable Energy Laboratory (NREL) have shown that dilute acid pretreatment is an effective, low cost, energy efficient and scalable means to recover lipids from wet microbial biomass (3134).

It is also necessary to consider the compatibility of the cell lysis method with lipid extraction. For example, hexane is a commonly employed extraction solvent, but does not always maximise extraction yields. In many cases, modification of solvent polarity (for example employing a mix of hexane with ethanol or isopropanol) is necessary for high lipid recovery (3539). However, incorporation of a polar solvent also often leads to more prevalent co-extraction of other polar materials, such as polar lipids, pigments, proteins and sterols, that require more severe cleanup of the lipid phase. In particular, extraction methods that incorporate chlorinated solvents, such as the well-known Folch and Bligh-Dyer extraction protocols, tend to extract significant amounts of polar lipids and other polar materials (36, 38). Combinations of hexane with alcohols, such as ethanol and isopropanol, have the potential to more selectively extract hydrophobic lipids while maintaining high extraction yields of these components (31, 39, 40). The optimal balance is thus a function of lipid speciation, content and speciation of other potential co-extractives, the lipid cleanup strategy and the hydroprocessing approach.

Lipid Composition

SCOs that are considered for biofuel production are mainly recovered from oleaginous algae, fungi (yeast and filamentous) and bacteria, with the number of known oleaginous species decreasing by family in the order listed. These microbes may be cultivated autotrophically (using only CO2 as a carbon source), mixotrophically (using a mixture of CO2 and organic carbon sources) or heterotrophically (using organic carbon sources, such as glucose and acetate). Several recent reviews have compiled oil content and broad composition of oleaginous microbes and their production of oils with an emphasis on wastes and lignocellulosic carbon sources (79, 11). A sampling of these compositions is reproduced in Table I, with comparison to common vegetable oils. It is important to note that analytical methods vary greatly across the references cited in Table I. The lipid content data, especially, should be taken with a grain of salt.

Table I

Lipid Content and Speciation in Common Oleaginous Microbes and Terrestrial Plants

Species Lipid content (% dry weight) Fatty acids (% total lipid)
Reference
C12:0 C14:0 C16:0 C16:1 C17:0 C18:0 C18:1 C18:2 C18:3 C20:0 C22:0
        Microalgae and cyanobacteria
Chlorella sp. 28–32 7–19 10–11 1–4 8–9 1–14 16–19 (41)
Chlorella zofingiensis 28–32 23 2 2 36 18 8 (41)
Crypthecodinium cohnii 23 13 23 3 8 (41)
Chatoceros muelleri 31–68 18–40 5–40 0.0–25.0 0–4 0–5 0–5 (41)
Schizochytrium linacinum 50–77 3–20 54–60 1–4 (41)
Various (10 species) 0.2–7.9 6.2–35.8 0.3–22.0 0.3–8.6 1.3–19.0 12.6–75.2 1.3–14.6 1.5–33.0 0.0–3.6 0.0–3.9 (42)
Dunalliela salina 6–27 6.9 2.8 9.8 14.7 3.8 8.0 4.9 12.9 1.7 24.8 (43, 44)
Synechocystis sp. PCC6803 18 19.9 20.9 43.3 11.3 1.5 1.1 1.4 (45)
        Yeast
Cryptococcus albidus 60 12 1 3 73 12 (41)
Cryptococcus curvatus 55–60 18.0–30.0 10.0–12.0 35–48 10–28 (10)
Lipomyces starkeyi 63 34 6 5 51 3 (41)
Rhodosporidium toruloides 66 18 3 3 66 (41)
Rhodosporidium toruloides 55–60 23–25 10–12 48–53 10–15 (10)
Rhodotorula glutinis 72 37 1 3 47 8 (41)
Trichosporon guehoae 65–78 20–22 10–25 35–38 12–22 (10)
Saccharomyces cerevisiae 30–50 8–10 32–43 4–7 42–48 (46)
Yarrowia lipolytica 36 11 6 1 28 51 (41)
        Bacteria
Acinetobacter baylyi 12.4 5 1 30 40 (47)
Alcanivorax borkumensis 23.2–26.9 2.9–15.0 36.8–85.3 tr–19.7 2.4–8.3 tr–41.2 (48)
Rhodococcus opacus 27.5–68.1 2.7–5.1 23.0–31.0 9.9–11.7 17.2–18.3 5.6–7.1 18.9–24.4 (49)
Rhodococcus opacus 19.0–26.0 3.0–19.0 6.0–74.0 (41)
Streptomyces coelicolor 64.2–83.0 8–27 28–38 1–3 1–3 (50)
        Filamentous fungi
Aspergillus sp. 12.4–22.1 0.8 14.4–14.8 22.5–29.6 43.4–48.7 0.4–1.6 0.4 0.1 3.0–12.2 0.1–2.9 (51)
C. echinulata 6.5–46.6 11.0–23.8 (52)
C. echinulata 5.3–8.4 22.5–24.7 6.2–8.2 35.3–38.5 18.3–20.5 12.0–13.8 (53)
M. isabellina 8.3–25.3 18.6–25.3 tr–4.9 2.0–6.2 45.9–50.4 13.6–20.6 3.5–5.5 (54)
M. isabellina 40–8–41.8 21.1–25.3 2.9–3.7 11.9–12.3 51.6–52.6 6.4–7.8 (55)
M. isabellina 34.0–50.0 20.3–29.0 2.0–2.7 3.0–5.0 41.2–55.0 3.0–27.8 (41, 56)
M. hiemalis 16.8–20.5 20.0–23.4 6.4–8.4 23.2–25.6 16.0–18.8 23.3–23.5 (53)
Mucor sp. 25.0–50.8 22.0–28.7 1.2–5.0 38.0–45.6 7.6–10.0 7.8–10.4 (41, 53)
Mucor sp. 1–9.4 13.8–24 tr–11.5 1.3–6.9 32.0–41.5 11.0–27.3 8.0–18.6 (54)
R. stolonifer 9.3–16.3 27.4–29.6 5.3–8.1 27.3–29.5 20.0–23.2 13.7–15.9 (53)
Rhizopus arrhizus 57 18 6 22 10 12 (41)
T. elegans 1–3.3 12.6–18.7 tr–2.3 4.8–12.4 38.7–73.3 5.6–15.7 2.4–23.0 (54)
Z. moelleri 18.1–22.0 27.3–29.5 14.4–16.6 23.5–25.9 12.8–14.2 13.3–15.5 (53)
        Plants
Brassica napus (rapeseed) 45 4 2 62 22 10 (41)
Elaeis guineensis (palm) 50 1 44 4 38 10 1 (41)
Glycine max (soybean) 8.1–27.9 10.0–11.0 4.0 18.0–24.0 54.0–55.0 7.0–13.0 (41, 57, 58)
Zea mays (corn) 3.0–11.0 0.0–0.3 8.6–16.5 0.0–0.05 0.0–3.3 20.0–42.2 34.0–65.6 0.0–2.0 (59)

Both SCOs and terrestrial plant oils typically contain high proportions of palmitic (16:0), palmitoleic (16:1), stearic (18:0), oleic (18:1) and linoleic (18:2) acids. On the surface, the similar composition of both oils appears to make SCOs an attractive feedstock for catalytic upgrading. However, SCOs can contain a broader distribution of lipid classes than terrestrial plant oils, including higher proportions of polar lipids, free fatty acids (FFA), sterols, terpenes, carotenoids and chlorophyll, as well as unidentified compounds, that can be co-extracted with the desired monoacylglycerides (MAG), diacylglycerides (DAG), triacylglycerides (TAG) and FFA. Examples of these other lipid classes are shown in Figure 1. The proportions of each vary greatly depending on the organism, growth mode, growth condition, growth stage and postharvest processing conditions, including biomass and extracted oil storage and solvent extraction conditions (36, 60). For example, glycolipids and phospholipids can account for 17–90% of the total lipids in autotrophically-grown algae (61), compared to ~2% in soybean oil (62), and lipase enzymes present in the cells can convert acylglycerides to FFA during lipid or biomass storage (60).

Fig. 1.

Lipid classes present in single-cell oils

Lipid classes present in single-cell oils

TAGs and FFAs are usually considered as the favoured precursors for biodiesel or hydrocarbon-based biofuels such as renewable diesel or renewable jet fuel. Other components are often detrimental in catalytic hydroprocessing as they either increase hydrogen consumption due to a high degree of unsaturation (for example terpenes, carotenoids and chlorophyll), contain heteroatoms that can poison metal or acid catalysts (for example polar lipids and chlorophyll) or contribute to poor cold flow properties (for example sterol-derived cycloparaffins). For example, polar lipids mainly consist of phospholipids, glycolipids, lipoproteins and sulfolipids. A phospholipid molecule consists of a polar phosphorus-containing moiety (such as phosphate, phosphatidylethanolamine or phosphatidylcholine), whereas a glycolipid contains a polar carbohydrate moiety in place of the phosphor group. The sidechains of polar lipids (sugars, proteins or phosphorous-containing molecules) may also include other nitrogen, sulfur or phosphorus-containing moieties (21).

On the other hand, hydrotreating of some of these contaminants can be beneficial in that the high degree of branching (for example in terpenes and the phytol chain of chlorophyll) can potentially improve cold flow properties and increase the value of the renewable diesel fuel. Additionally, when a large proportion of the lipid fraction comprises nominally undesirable molecules (such as polar lipids), removal of the entire molecule in a cleanup strategy would result in low fuel yields. Thus, while some cleanup of the oils is likely necessary, the most desirable cleanup strategy is not universal across SCOs, and tradeoffs between overall yields and catalyst longevity must be considered.

Lipid Cleanup

Cleanup of SCOs targets many of the same impurities that are removed in refining of vegetable oils for human consumption, and thus cleanup approaches can be broadly categorised into the same four operations: bleaching to remove pigments and polar compounds, degumming to precipitate phospholipids, deodorisation to remove FFAs and distillation to remove additional FFAs and other volatile matter. When refining SCOs for hydrotreating, some modifications may be necessary. For example, FFAs are one of the preferred feeds for hydrotreating so deodorisation and distillation will not always be needed. However, in some cases it may be advantageous to hydrolyse the entire lipid stream to FFAs, distil these FFAs away from impurities, and route the distillate to hydrotreating. In these cases, bleaching and degumming may not be needed. Researchers have explored to some extent each of these steps except deodorisation. These techniques and their advantages and disadvantages are summarised in Table II.

Table II

Strategies for Cleaning Up Single Cell Oils

Cleanup technology Advantages Disadvantages
Adsorbent bleaching Relatively inexpensive May not fully remove impurities
Established technology May retain oil on solid, decreasing yields
Suitable for broad range of impurities May retain polar lipids, decreasing yields significantly for oils high in these components
Acid bleaching May be more effective than adsorbents for chlorophyll Requires additional water to remove acid from oil
Targets chlorophyll, may not be effective for other impurities
Degumming Relatively inexpensive Targets polar lipids, decreasing yields significantly for oils high in these components
Established technology May not remove polar lipids that are non-hydratable
May be simple to integrate with cell lysis by dilute acid May not remove impurities other than polar lipids
Hydrolysis and FFA distillation Suitable for broad range of impurities and lipid compositions Relatively expensive for cost of alkali
Relatively energy intensive
(Trans)esterification and fatty acid methyl ester (FAME) distillation Suitable for broad range of impurities and lipid compositions Relatively expensive for cost of alkali, acid or methanol
Relatively energy intensive

Bleaching

Crude extracted oils from autotrophically-grown microbes are often high in chlorophyll content. The removal of chlorophyll is important in both hydroprocessing and biodiesel production as chlorophyll contamination can deactivate catalysts and degrade fuel quality (60, 63, 64). For example, the porphyrin head of the chlorophyll molecule is highly unsaturated and contains the catalyst-poisoning heteroatoms nitrogen and magnesium. The two most common approaches to remove chlorophyll are adsorption on a solid material and dealkylation via acid treatment.

Bleaching earths, activated carbon and silica are common adsorbents for oil refining. Adsorption of chlorophyll is initiated by ion exchange of the Mg2+ centre of the porphyrin head with a proton. The metal-free porphyrin is then protonated and adsorbed (65). Adsorbents are commonly pretreated with mineral acids to increase the number of exchangeable protons (66). Ultrasound-assisted adsorption has also been demonstrated to increase the rate of chlorophyll removal from oils (67). It is worth noting that the loadings of polar adsorbents to fully remove chlorophyll from SCOs may be much higher than typically used for edible oil bleaching due to the higher levels of chlorophyll (and other adsorbing contaminants). For instance, Chen et al. employed bleaching earth to adsorb chlorophyll from microalgal biodiesel at a mass loading of 16 wt% (60), whereas a typical edible oil bleaching process uses a mass loading of 1.5 wt% (68). Increasing adsorbent loading, while effective in reducing chlorophyll content, decreases yields due to co-adsorption of lipids. This is particularly true of bleaching earths, which can retain up to a third of their mass in oil (69), though even higher retentions have been claimed (70). Similarly, Santillan-Jimenez et al. removed chlorophyll from crude extracted algae oil via adsorption on K10 montmorillonite clay and activated carbon, recovering only 58 wt% and 46 wt% of the original extract mass, respectively (71). Gas chromatography-mass spectrometry (GCMS) analysis of the recovered materials indicated a high content of fatty acids, hydrocarbons and phytol isomers, while ultraviolet-visible (UV-vis) spectroscopic and inductively coupled plasma (ICP) analyses revealed complete chlorophyll, phosphorus and magnesium removal. Notably, the relative abundance of hydrocarbons in the activated carbon-bleached material was lower than in the K10-bleached material, likely due to the greater hydrophobicity of the activated carbon retaining more of the hydrocarbons. This observation is also consistent with the lower total recovered yield from the carbon adsorbent. Thus, adsorbents with higher polarity may be more selective for removal of the target compounds.

Phosphoric acid-catalysed dealkylation was originally developed by Diosady to refine Canadian canola oil, which has anomalously high chlorophyll content for a terrestrial oil due to weather conditions in the region (70). The acid bleaching approach employs anhydrous acids, such as phosphoric acid or sulfuric acid to cleave the phytol side chain from the porphyrin structure, the latter of which is not soluble in oil. Thus, this precipitable form of pheophorbide can be easily separated from the oil by filtration while the hydrophobic phytol is preserved. Retaining the phytol in the oil confers the additional benefit of improved biofuel yield and performance due to a high degree of branching. Dong et al. employed a modified version of Diosady’s method to remove 99.7% of chlorophyll from algae oil (63). In a direct comparison of adsorptive and dealkylative approaches, Kruger et al. found that phosphoric-acid-catalysed dealkylation was more effective than adsorption on silica for the removal of chlorophyll from algae oil (72). In the comparison, adsorption on silica gel removed 85% of the oil nitrogen content and still showed a detectable chlorophyll signal in UV-vis analysis, while acid bleaching removed 92% of the oil nitrogen and did not show a detectable chlorophyll UV-vis signal. Moreover, phosphoric acid bleaching is compatible with conventional vegetable oil refining practice, since phosphoric acid is routinely added for oil degumming. However, for the same reason and also because the acid must be washed out of the oil by adding water, phosphoric acid bleaching may suffer from yield losses and poor bleaching performance in oils with high levels of phospholipids.

Apart from adsorption and acid treatment, other novel methods of chlorophyll removal have been reported. Sathish developed a multi-step process to remove chlorophyll from microalgal crude oil, involving sequential aqueous acid, base and acid treatments to lyse cells, saponify lipids and precipitate chlorophyll, respectively (73). Though the extraction process was not optimised, roughly 20% of the lipids remained with the biomass solids and another 20% with the precipitated chlorophyll. However, the purified lipids showed no chlorophyll signal in UV-vis analysis. The process is advantageous in that it does not require anhydrous conditions, but the sequential use of concentrated acids and bases is likely to be expensive and generate highly saline wastewater. On the other hand, Li et al. removed chlorophyll from intact microalgae by saponification with sodium hydroxide (74).

Degumming

Analogous to chlorophyll, SCOs can have higher phospholipid content than most terrestrial oils, and removal of these polar lipids is necessary to prevent catalyst fouling in downstream processes. Further complexity arises as a large portion of phospholipids in SCOs are non-hydratable, namely phosphatidic acid and phosphatidylethanolamine (75). These species complex with metal cations and cannot be removed by conventional water degumming processes that precipitate hydratable phospholipids. Other degumming methods have been developed to remove these non-hydratable phospholipids (76).

Acid degumming is the most common approach to removing non-hydratable phospholipids. An acid is added to decompose phospholipid salts to improve their hydratability. Phosphoric acid is routinely used in algae oil degumming given the co-benefit of chlorophyll reduction (32, 54, 60). A drawback of acid degumming, however, is that some decomposed phosphatic acid will remain in the oil phase (76). Subsequent addition of a small amount of diluted base – enough to neutralise phosphatic acid while avoiding saponification – has been shown to be effective in further reducing phosphorus content of acid-degummed oils (77). Other methods which target both hydratable and non-hydratable phospholipids, such as membrane or enzymatic degumming, are in nascent stages of development. While promising, the cost of these techniques are prohibitively expensive for SCO degumming in their current state (76).

Given the potentially high content of microbial polar lipids, a remarkable amount of microbial lipid may be lost in the degumming process (60). Nevertheless, in some SCOs degumming may be appropriate, and examples of both dilute acid and solvent degumming exist in the patent literature (7880). For SCO streams containing large fractions of polar lipids, hydrolysis to a FFA stream is likely preferable for fuel production (34).

Hydrolysis and Distillation

As an alternative to removing impurities from the lipid stream, approaches have also been developed to remove only the desired components from the lipid stream and leave everything else. Within this realm, hydrolysis and methanolysis are the primary approaches applied, generating FFA and FAME, respectively, both of which are distillable.

Hydrolysis can employ acids, bases or high temperature water to cleave fatty acid and other esters. Alkaline hydrolysis (i.e. saponification) is advantageous in that it requires mild temperatures and short reaction times (typically 80°C and 1 h, respectively). However, the stoichiometric consumption of alkali likely makes this approach too expensive for a fuel production scenario. Acid hydrolysis is favourable in that the acid is catalytic and mild temperatures are also typically employed (100°C or less), but reaction times are typically 8–24 h. Acid hydrolysis was the basis of the industrial Twitchell process, which was subsequently superseded by steam splitting. Steam splitting via the Colgate-Emery process is the current industrial standard, employing temperatures of 250–330°C and reaction times of 2–3 h (81). Higher temperatures can significantly shorten the reaction time (for example to ~10 min), but also significantly increase the operating pressure (for example to ~2500 psig) (82). Steam splitting is effective with both acylglycerides and with phospholipids (82). Lawal et al. demonstrated the effectiveness of hydrolysing algal lipids to FFAs and distilling the hydrolysed FFAs away from impurities (83). An alternative approach involves engineering microbes to produce and excrete FFAs directly into the growth medium, which could simplify the cell lysis and lipid extraction operations. This approach was recently demonstrated in the cyanobacteria Synechocystis sp. PCC6803 (45).

Alternatively, microbial lipids can be converted to methyl esters, which facilitates distillation without vacuum. The (trans)esterification to methyl esters is widely used on a commercial scale to produce biodiesel, and occurs under similarly mild conditions as saponification. Murzin and coworkers demonstrated the effectiveness of this cleanup approach for removing impurities in algae oil (8486). However, this approach suffers from two significant drawbacks, namely that it requires a dry feedstock and that it consumes methanol stoichiometrically. Drying of microbial biomass is much more energy intensive than drying terrestrial crops, and may be economically untenable for fuel production. Process engineering may make it feasible to recycle the methanol (or valorise it, for example in a methanol-to-gasoline-type process) though this has not been explored to our knowledge.

Hydroprocessing of Single Cell Oils

Several types of SCOs have been upgraded to hydrocarbon fuels with and without prior cleanup. Among these, algae oils are by far the most common. Hydroprocessing approaches and cleanup approaches for use on algae oils have varied significantly, though many employed techniques to specifically remove chlorophyll. Hydroprocessing of yeast oils, which do not contain chlorophyll but may contain sterols, has also been reported by a few researchers. Among other microbes, to our knowledge only lipids from the methanotrophic bacterium Methylomicrobium buryatense have been upgraded to hydrocarbons. A summary of SCO hydroprocessing literature is provided in Table III.

Table III

Summary of Single Cell Oil Hydroprocessing Reports

Reference SCO type Oil recovery Oil cleanup Deoxygenation
Hydroisomerisation
Conditions Catalyst Results Conditions Catalyst Results
(87) Algae, Nannochloropsis salina Obtained from Valicor 1 mm filtration 1.3 wt% oil in n-C12H24, 280–360°C, 300–500 psig H2, flow NiMo/Al2O3 63 wt% hydrocarbon yield N/A N/A N/A
(88) Algae, Nannochloropsis salina Obtained from Valicor 1 mm filtration 1.3 wt% oil in n-C12H24, 280–360°C, 300–500 psig H2, flow Pt/Al2O3, Rh/Al2O3, NiMo/Al2O3 76 wt% hydrocarbon yield (Pt/Al2O3), 56% hydrocarbon yield (Rh/Al2O3), 63 wt% hydrocarbon yield (NiMo/Al2O3) N/A N/A N/A
(71) Algae, Scenedesmus acutus Modified Bligh-Dyer extraction K10 montmorillonite clay or activated carbon in column chromatography 1.3 wt% oil in n-C12H24, 260–300°C 580 psig H2, flow Ni-Al layered double hydroxide Hydrocarbon yields not quantified, but GCMS suggested nearly complete conversion to hydrocarbons at 1 h TOS, decreasing to 65–70% at 4 h TOS at 260°C, while conversion remained nearly complete through 4 h TOS at 300°C N/A N/A N/A
(89) Algae, Scenedesmus acutus Modified Bligh-Dyer extraction Activated carbon, silica gel in column chromatography 1.3 wt% oil in dodecane, 260°C, 580 psig H2, flow Ni/Al2O3, Ni-Cu/Al2O3 Hydrocarbon yields not quantified, but GCMS suggested incomplete conversion to hydrocarbon even at 1 h TOS. Catalyst deactivated faster with algae oil than with triolein N/A N/A N/A
(90) Algae, Scenedesmus acutus Modified Bligh-Dyer extraction K10 montmorillonite clay in column chromatography 1.3 wt% oil in dodecane, 300°C, 580 psig H2, flow Ni-Al layered double hydroxide 95% conversion, 73% selectivity to C10–C17 N/A N/A N/A
(91) Algae, Scenedesmus acutus Modified Bligh-Dyer extraction K10 montmorillonite clay in column chromatography 1.3 wt% oil in dodecane, 260°C, 580 psig H2, flow Ni-Al layered double hydroxide 55–65% hydrocarbon in product mixture, lower hydrocarbon yield but increasing diesel-range selectivity with increasing TOS N/A N/A N/A
(92) Algae, not specified Obtained from Verfahrenstechnik Schwedt GmbH None 1.3 wt% oil in dodecane, 260°C, 580 psig H2, flow Ni/HBEA zeolite 100% conversion, 78% hydrocarbon yield, 60% n-C18 yield N/A N/A Noted 3.9–11% isooctadecane yield in batch reactions
(93) Algae, not specified Obtained from Verfahrenstechnik Schwedt GmbH None 1.3 wt% oil in dodecane, 270°C, 580 psig H2, flow Ni/ZrO2 100% conversion, 75% hydrocarbon yield, 70% n-C17 yield N/A N/A Noted modest isomerisation activity, but did not report isomer yields
(94) Algae, not specified Obtained from Verfahrenstechnik Schwedt GmbH None 1 g oil in 100 ml dodecane, 260°C, 580 psig H2, batch Ni/HBEA zeolite 100% conversion, 79% hydrocarbon yield, 40–43% n-C18 yield, 25–27% n-C17 yield N/A N/A 2–3% isooctadecane yield
(95) Algae, Chlorella sp. Supercritical hexane None 1 g oil in 100 ml dodecane, 300°C, 440 psig H2, batch Ni/HY zeolite, Ni/SiO2, Mo2N/MoO2 32–49% FFA yield, 4–7% hydrocarbon yield N/A N/A N/A
(96) Algae, Chlorella sp. Supercritical hexane None 1 g oil in 100 ml dodecane, 300°C, 440 psig H2, batch Ni/SiO2 15% conversion, 6% hydrocarbon yield, 42% FFA yield N/A N/A N/A
(72) Algae, Scenedesmus acutus Acid pretreatment, hexane liquid liquid extraction Silica gel in column chromatography, H3PO4 bleaching 25 wt% oil in hexane, 450°C, 1300 psig H2, flow Pd/C 73–74% liquid hydrocarbon yield, 15–17% C4–C11, 82–85% C12+ Neat hydrodeoxygenation product, 350°C, 500 psig H2 Pt/SAPO-11 77–81% liquid hydrocarbon yield, 30–50% isomer content, –12 ≤ CP ≤ –21°C
(97) Algae, Scenedesmus acutus Acid pretreatment, hexane liquid liquid extraction Silica gel in column chromatography, H3PO4 bleaching 25 wt% oil in hexane, 450°C, 1300 psig H2, flow Pd/C 69% liquid hydrocarbon yield, 21% C4–C11, 79% C12+ Neat hydrodeoxygenation product, 350°C, 500 psig H2 Pt/SAPO-11 69% liquid hydrocarbon yield, 32% isoparaffin content, cloud point = –3.5°C
(98) Algae, not specified Obtained from Phycal None Neat oil, 350°C, 800 psig H2, flow Pd/C, Pt/Al2O3 85% of liquid product stream as n-alkanes (95% with second pass), polishing with Pt/Al2O3 converted residual oxygenates Neat hydrodeoxygenation product, 350°C, 800 psig H2 Pt/USY 50% isomerisation conversion, solvent dewaxing resulted in product that remained liquid at –20°C
(99) Algae, not specified Obtained from Xian Lyphar Biotech Co Ltd None Neat oil or 5 vol% oil in heptane, 260–300°C, 580 psig H2, batch Co/natural clay 84–100% conversion of oil, 83–86% yield of alkanes N/A N/A N/A
(100) Algae, Chlorella Extraction with CH2Cl2 None Oil in H2O (ratio unspecified), 330–370°C, batch Pt/C Yields not quantified, but increase in heating value and carbon content of oil phase, and highly isomerised products identified in liquid product N/A N/A N/A
(101) Algae, not specified Obtained from Solix Biofuels None 10 mg oil in 10 ml hexanes, 290°C, 440 psig H2, batch Fe/MSN 67% conversion, 16% alcohols, 33% unsaturated hydrocarbons, 18% saturated hydrocarbons, remaining FFAs all saturated N/A N/A N/A
(102) Algae, Scenedesmus obliquus Extraction with CHCl2:methanol None Neat oil, 290–350°C, 60 bar H2, batch NiMo/Al2O3 84–87% liquid yields, increasing selectivity toward alkanes at higher temperature and catalyst loading N/A N/A N/A
(103) Algae, Nannochloropsis sp. Incubation, high pressure homogenisation, hexane extraction None Neat oil, 300–375°C, 50–120 bar H2, batch CoMoP/Al2O3 >99% conversion, up to 80% liquid yield, giving way to cracking and polyaromatisation at higher pressures N/A N/A N/A
(104) Algae, Dunaliella sp. Electroflocculation and osmotic lysis, self-separation Hot water hydrolysis Oil in dodecane, 300°C, 19 bar 6% H2, fed batch Pd/C 98% conversion to n-alkanes N/A N/A N/A
(105) Cyanobacteria, Synechocystis PCC 6803 Excreted FFAs, recovered by solvent washing adsorbent Recrystallisation, activated carbon, saponification, reverse phase liquid chromatography 1.15 g oil in 22.5 g n-C12H24, 300°C, 15 bar of 5% H2, fed batch Pd/C 32–88% alkane yield 25 g n-alkane, 300°C, 500 psig H2 Pt/CaY zeolite 20–36% conversion, 18–31% yield to isoparaffins, 92–95% isomerisation selectivity
(10) Yeast, Rhodosporidium toruloides Acid pretreatment, hexane liquid liquid extraction None 25 wt% oil in hexane, 450°C, 1300 psig H2, flow Pd/C 80% liquid phase yield, 60% C12–C20 hydrocarbons, 12% C7–C11 hydrocarbons, 25% C20+ hydrocarbons Neat hydrodeoxygenation product, 350°C, 500 psig H2 Pt/SAPO-11 68% liquid hydrocarbon yield, 33% isoparaffin content, 70% C12–C20 hydrocarbon, 20% C7–C11 hydrocarbon, 7% C20+ hydrocarbon, cloud point –14.5°C (after cold filtration)
(34) Bacteria, Methylomicrobium buryatense Alkali-acid pretreatment, hexane extraction Alkali-acid pretreatment, hexane extraction 0.5 ml oil in 25 ml decane, 360°C, 40 bar H2, batch Pd/SiO2 100% conversion, ~100% yield to C13–C17 paraffins N/A N/A N/A

Lawal and coworkers explored deoxygenation of algae oils from Nannochloropsis salina and Chlorella vulgaris over platinum, rhodium and nickel-molybdenum-based catalysts (83, 87, 88). The oils were obtained from Valicor, which employed a thermochemical pretreatment of the algal biomass to render the lipids extractable (83, 106). Notably, this pretreatment reduced the amount of phosphorus and iron in the Chlorella oil by nearly 100%, while sulfur was only reduced by 30%. For the N. salina oil, however, the sulfur and phosphorus content remained relatively high at 2033 ppm and 246 ppm, respectively. Additionally, the N. salina oil contained only 47% material that could be identified as acylglycerides or FFAs (83, 87). Analysis showed that the oil had up to 0.8% chlorophyll, 0.5% carotenoids, 5% sterols, 1–5% mannitol and a large fraction of unsaponifiable matter that was not identified (83). While most reaction conditions explored showed significant catalyst deactivation within 7 h time on stream (TOS), Zhou and Lawal were able to find a set of conditions for both a NiMo/Al2O3 and a Pt/Al2O3 catalyst that did not show significant deactivation within this timeframe, and obtained hydrocarbon yields above 60 wt% (87, 88). The optimal conditions for the platinum and nickel-molybdenum catalyst were significantly different, underscoring the need for reaction engineering studies for each combination of catalyst and oil.

Crocker and co-workers explored deoxygenation of algae oils from Scenedesmus acutus grown on power plant flue gas as a carbon source using nickel-based catalysts (71, 8991). While the focus of these experiments was generally on model compounds, all of the catalysts employed with algae oils (Ni/Al2O3, Ni-Al layered double hydroxide and Ni-Cu/Al2O3) deactivated faster with the algae oils (within 4 h TOS) than with vegetable oils or model compounds such as triolein, despite the fact that the oil concentration was only 1.3 wt% in a dodecane solvent. The fast deactivation may have been in part due to the low temperature employed (260°C). In particular, the Ni-Al layered double hydroxide catalyst showed significantly less deactivation at 300°C. The faster deactivation with algae oils is also notable in light of the cleanup procedures employed: the lipids were purified by column chromatography using either K10 montmorillonite clay or activated carbon with silica gel, which was effective at removing chlorophyll and other pigments, magnesium and phosphorus. K10 montmorillonite appeared to be a more effective adsorbent than activated carbon, resulting in a greater fraction of products in the diesel range. The authors hypothesised that several unquantified components in the algae oil, including FFAs, polar lipids, sterols or fatty amides could be contributing to the deactivation (90). However, experiments with mixtures of model TAGs and FFAs suggested FFAs were not the culprit (90). A more detailed analysis of impurities also suggested that the oil cleanup quantitatively removed phosphorus and magnesium, though a small amount of nitrogen remained (71). Santillan-Jimenez et al. (71) also noted that the residual extraction solvent (namely chloroform) was likely the source of chloride detected on the catalyst in post-reaction analysis (71), underscoring the observation that the extraction protocol can impact downstream processes through routes other than influencing the types of lipids extracted.

Lercher and coworkers explored nickel-based catalysts for algae oil deoxygenation under similar conditions to those above (though also in batch mode), using commercial algae oil of an unspecified species (9294). In these experiments, support acidity (HBEA zeolite or zirconia) was shown in mechanistic studies with model compounds to play a key role in the reaction kinetics, and the nickel particle size was also key to catalyst activity. The oil was not analysed for impurities (only fatty acid composition), but did not deactivate the catalysts for at least 72 h (for Ni/ZrO2) (93) or 120 h (for Ni/HBEA) (92) TOS. Notably, some isoalkanes, which improve the cold flow properties of the diesel blendstock, were observed in the products due to the support acidity.

Nguyen et al. explored both nickel and molybdenum-based catalysts in deoxygenation of Chlorella algae oil extracted by supercritical hexane (95, 96). In these studies, the catalysts were rapidly deactivated (within 1 h) and produced less than 10% yield to hydrocarbons. In contrast, FFAs were the major product, with acylglycerides featured prominently as well. The authors noted that the crude extracted lipids comprised only around 50% fatty acids and 3–4% sterols, with the remainder unidentified (95). While low conversion of the FFA and acylglycerides was observed, no conversion of the sterols or steryl esters was found (96). Nguyen et al. also noted that while molybdenum nitride catalysts had much lower initial activity than nickel-based catalysts, the molybdenum nitride catalysts did not show as strong a deactivation as the nickel catalysts did (95). The low activity of the molybdenum nitride catalysts represents a significant challenge nonetheless. However, recovery of the lipids as methyl esters and purification over a bleaching clay produced a feedstock that could be readily converted over a Ni-HY zeolite catalyst (8486). Interestingly, a Pd/C catalyst did not perform as well as the Ni-HY catalyst in deoxygenation of the purified methyl esters, possibly due to the lower acidity of the carbon support.

Researchers at NREL demonstrated both deoxygenation and hydroisomerisation of Scenedesmus algae oil recovered after acid pretreatment and hexane extraction (72, 97). Deoxygenation employed a Pd/C catalyst, while hydroisomerisation employed a Pt/SAPO-11 catalyst. Kruger et al. found that while relatively severe temperature and pressure (450°C and 1300 psig) and dilution of the algae oil in a hexane solvent were required for complete conversion and stable performance of the Pd/C deoxygenation catalyst, bleaching the oils with either silica gel or with phosphoric acid did not improve the hydroisomerisation performance, at least through 10 h TOS (72). The deoxygenation step also resulted in significant denitrogenation as well. The severe conditions required for deoxygenation led to significant cracking and lower yields to diesel range products, but hydroisomerisation produced renewable diesel blendstocks with cloud points below –10°C (72). The latter step is important to demonstrate given the non-zero levels of residual nitrogen in the deoxygenated feedstock that could poison acidic isomerisation catalysts.

Robota et al. also employed a Pd/C catalyst for deoxygenation of a neat algae oil, and then hydroisomerised the resulting alkanes with a Pt/USY zeolite catalyst (98). Despite the higher concentration of oil, less severe conditions were required for nearly complete conversion of the oil to alkanes than was observed by Kruger et al. (72), though a second pass and a polishing step to remove residual oxygenates were used to produce a clean alkane stream for hydroisomerisation. With the large-pore Pt/USY zeolite catalyst, isomerisation and cracking were competitive, resulting in a 40% or more mass loss to naphtha-range hydrocarbons. Though cloud points of the resulting products were not measured, solvent dewaxing yielded a product that remained liquid at –20°C.

Soni et al. (99) evaluated cobalt/natural clay catalysts for deoxygenation of a commercial algae oil, and achieved hydrocarbon yields around 85% after 8 h in a batch reactor at 580 psig hydrogen. This oil contained nearly 70% docosahexaenoic acid in its FFA profile, suggesting that the product alkanes would require some degree of cracking to serve as a diesel blendstock.

Fu et al. (100) demonstrated hydrogen-free decarboxylation of Chlorella oil at 330–370°C for 2 h in a batch reactor. The heating value and carbon content of the organic phase both increased and the organic product contained significantly isomerised compounds, though conversion, yield, oil characterisation and catalyst characterisation were not reported.

Kandel et al. (101) reported the conversion of an algae oil from Solix Biofuels into hydrocarbons using a nanoparticle iron catalyst supported on mesoporous silica. After 6 h at 290°C and 440 psig hydrogen, 67% of the algae oil had been converted to both saturated and unsaturated alcohols and hydrocarbons. The authors noted that the presence of unsaturated compounds suggests hydrogen may have been limiting.

Tang et al. (102) evaluated deoxygenation of a Scenedesmus algae oil in a batch reactor over a NiMo/Al2O3 catalyst. As the reaction temperature increased from 290°C to 350°C (and as catalyst loading increased from 0 wt% to 50 wt% of the oil), the product spectrum shifted from acids, esters and alcohols to paraffins, olefins, naphthenes and aromatics, with a small amount of isoparaffins also present. The authors used multiple solvent combinations to extract the lipids. Dichloromethane:methanol was the highest yielding solvent on a weight basis, but the primary components were sterols and tocopherols rather than acylglycerides and FFAs. Other solvents were lower yielding, but the extracts were also not analysed.

Poddar et al. deoxygenated Nannochloropsis lipids over a phosphorus-promoted CoMo/Al2O3 catalyst, achieving nearly complete conversion to alkanes and nearly 80% yield to liquid-phase products in 6 h at 375°C and 50 bar hydrogen (103). The authors conducted a detailed kinetic analysis but did not report detailed product analysis or post-reaction catalyst characterisation.

Wang et al. explored deoxygenation of a FFA stream derived from Dunaliella lipids (104). The lipids were hydrolysed in 250°C to release FFAs, which self-separated from the aqueous phase and were decarboxylated in fed-batch mode over a Pd/C catalyst. The catalyst maintained steady operation over 5 h of reaction, even with a hydrogen-poor headspace in the reactor.

Finally, Schulz et al. (105) deoxygenated Synechocystis (cyanobacteria) lipids over a Pd/C catalyst and isomerised the resulting alkanes over a Pt/CaY zeolite catalyst. The lipids were unique in that they were recovered in the form of excreted FFAs (mainly C12), adsorbed on and eluted from a resin. Once recovered, the lipids were also subjected to a number of cleanup protocols, including recrystallisation, activated carbon treatment, saponification and preparatory-scale liquid chromatography. These protocols had varying effectiveness in reducing sulfur and phosphorus contaminants in the oils (especially sulfur), which were reflected in varying performance of the Pd/C deoxygenation catalyst across FFA streams, giving 33–88% alkane yield depending on the stream. The two most promising streams gave relatively low conversion, but high isomerisation selectivity over the Pt/CaY catalyst.

Aside from algae lipids, yeast lipids have also been evaluated for deoxygenation and hydroisomerisation. Sànchez i Nogué et al. (10) employed the same conditions as those used by Kruger et al. (72) and Knoshaug et al. (97) to deoxgygenate lipids from Rhodosporidium toruloides yeast. The yeast lipids produced a renewable diesel blendstock of similar quality as that from the algae oils. Process optimisation to determine if the yeast lipids could be satisfactorily converted under less severe conditions due to the lack of impurities such as chlorophyll was not conducted. The lipids were found to contain a small amount of steryl esters and a significant amount of polar lipids as well.

Chuck and coworkers have also evaluated conversion of yeast lipids to fuels, though not through conventional hydroprocessing approaches (107, 108). In these studies, the yeast lipids were either catalytically cracked (107) or metathesised with ethylene using a Hoveyda-Grubbs catalyst (108). Although the Pd/C cracking catalyst and reaction temperatures were similar to those used for deoxygenation by other researchers, the gas atmosphere was quite different. Interestingly, however, the catalytic cracking experiments were performed with a yeast lipid from Metschkownia pulcherrima that included 9 wt% sterols, and the authors inferred that the sterols can act as hydrogen donors as they undergo aromatisation at 350–400°C in an argon atmosphere, which improved the yield to linear alkanes from the triglyceride fraction.

Among other oleaginous microbes, the only report of hydroprocessing to our knowledge is that of Dong et al. (34), who converted polar lipids from Methylomicrobium buryatense to linear alkanes over a Pd/SiO2 catalyst. The bacterial biomass was hydrolysed by a sequential alkaline-acid method that both ruptured the cell wall (rendering the lipids extractable) and cleaved the polar head groups off the predominantly polar lipids, producing a clean FFA stream for deoxygenation. The high degree of polar head group cleavage was confirmed by the presence of only 4 ppm phosphorus in the oil, though the oil contained sulfur, nitrogen, sodium and halogens in higher amounts. With a relatively high catalyst loading and deoxygenation in batch mode, full conversion to linear alkanes was achieved. The authors also noted that a preliminary catalyst screening with nickel- and copper-based catalysts confirmed the hypothesis that these catalysts would be rapidly deactivated in the presence of phosphorus-containing feeds, while noble metals performed better.

Summary and Outlook

Several interesting themes emerge in lipid composition, cleanup and hydroprocessing of microbially-derived oils. First, the studies reporting the longest and most stable catalytic performance have used refined algae oils, for example from Phycal or Solix. Although the details of the refining process are not readily available, these processes appear to sufficiently clean the oils of catalyst-deactivating impurities. In contrast, lipid streams extracted by researchers directly prior to catalytic processing have tended to deactivate catalysts more rapidly or perform more poorly, even when diluted by a factor of 100 into an inert solvent. Perhaps this is unsurprising given the differing descriptions of the oils’ appearances, for example bright orange transparent liquid vs. dark brown semisolid. The situation is additionally confounded by the variety of techniques and solvents used to extract the lipids, which may co-extract different undesirable components. Hexane is the simplest solvent employed, but can result in low extraction yields and form emulsions with wet biomass. Alcohols, such as methanol and ethanol can be added to increase the extraction yields and minimise emulsions, but can also increase co-extraction of impurities. Halogenated solvents can serve the same purpose as alcohols, but further confound the process by introducing additional heteroatoms (typically chloride from chloroform or dichloromethane) that can remain in the extract in trace amounts even after solvent removal.

Second, studies using noble metal catalysts have tended to report better performance than those using base metal catalysts. This is frequently true in other catalysis applications as well, wherein base metals such as nickel, molybdenum and copper are more prone to coking, oxidation by feedstock oxygen and poisoning by feedstock heteroatom impurities. Indeed, in the examples where microbial oils have been thoroughly characterised for heteroatom impurities, it is tempting to infer that supported noble metals may even be able to tolerate some degree of phosphorus, nitrogen and sulfur in the feedstock. However, times on stream in the literature reviewed here, less than 200 h TOS, have typically been too short to make a meaningful assessment. Nevertheless, while noble metals are frequently avoided due to the high cost, there may be environmental advantages (at least for metals such as ruthenium and platinum) (109), and some of the cost can be recouped by regeneration and recycling of the spent catalysts (110). It is worth noting that industrial processes producing green diesel fuels from vegetable oils, such as Honeywell-UOP’s Ecofining, Neste’s NExBTL, Axens’ Vegan and Renewable Energy Group’s Bio-Synfining processes employ nickel-molybdenum or cobalt-molybdenum sulfide catalysts similar to those used in petroleum refining (111). These catalysts tend to favour hydrodeoxygenation over decarbonylation or decarboxylation, which preserves carbon in the product, but also consumes more hydrogen in the process. At least one evaluation of the tradeoffs between carbon yield and hydrogen consumption concluded that decarboxylation would be economically favourable, provided that hydrogen consumption through subsequent methanation and reverse water-gas shift reactions is not significant (112). On the other hand, the deployment of large-scale wind and solar power has at times resulted in excess power production and low-cost electricity that can be used for water electrolysis to produce inexpensive hydrogen. Nickel-molybdenum catalysts are also frequently employed for hydrodesulfurisation and hydrodenitrogenation, which may indicate a higher level of robustness to heteroatoms than other types of catalysts. In particular, nickel-molybdenum catalysts typically require a co-fed source of sulfur, often hydrogen sulfide or dimethyl disulfide, to mitigate conversion of the active sulfide phase to a less-active oxide. Concomitantly, these catalysts tend to leach sulfur into the product (21), which may inhibit the ability of these catalysts to meet increasingly stringent sulfur limits in fuels. It is unknown whether the sulfur-containing species in microbial oils can satisfy this requirement of nickel-molybdenum catalysts.

Third, lipid stream cleanup by a number of approaches appears promising, but all of the techniques have some drawbacks and only a few can be reasonably expected to apply universally across microbial lipids. Saponification of the lipid stream to FFA followed by distillation is perhaps the most robust as it should be effective even with lipid feeds containing high levels of polar lipids, sterols, chlorophyll and metals. However, the distillation step is energy intensive compared to most of the other bleaching methods and the cost of the alkali is high. Methyl esterification and distillation should be similarly effective, but the methanol would need to be recovered for high material and economic efficiency, or else recovered as methane after hydrotreating. Degumming (precipitation of phospholipids) and phosphoric acid bleaching can work well for lipid streams that are low in polar lipids, but would suffer from significant yield losses otherwise, and the phosphoric acid must be thoroughly washed out of the cleaned lipid stream to avoid contamination from the bleaching agent. In lipid streams with high polar lipid content, hydrolysis to FFAs may prove more economical. Adsorption on a polar material such as silica, bleaching clay or activated carbon appear effective for some target impurities (for example chlorophyll and some metals), but similar to degumming, may result in unacceptable yield losses for lipid streams that contain large amounts of polar lipids. Additionally, post-adsorption oils have still tended to deactivate the catalysts. Interestingly, even when chlorophyll is removed to below detection limit, some nitrogen frequently remains in the oils, likely in the form of hydrophobic protein. There is thus a clear need for both fundamental science and process development on this topic.

Looking forward, converting microbial lipids to fuels is a promising approach to displace conventional fossil fuels, especially for diesel and jet fuels which share the same carbon number range as the microbial lipids. While previous research has focused primarily on algae lipids, the advent of third generation cellulosic sugars may spur further development of heterotrophic oleaginous microbes as well. Finally, to best allow for economically-viable biofuels production from SCOs, it is critical to design an integrated process, in which certain complementary units can be intensified to maximise the yield of high quality products for a given step. For example, the lipid hydrolysis step can be integrated with the biomass pretreatment or cell wall rupture step to simultaneously rupture the cell walls for bulk oil extraction and liberate FFA from polar lipids. In addition, the selectivity of the extraction solvent should be carefully considered to increase extraction of fatty acids while reducing the co-extraction of impurities. With these considerations, SCOs are poised to make a significant impact in sustainable fuel production.

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References

  1. 1.
    E. J. Martínez, V. Raghavan, F. González-Andrés and X. Gómez, Int. J. Mol. Sci., 2015, 16, (5), 9385 LINK https://doi.org/10.3390/ijms16059385
  2. 2.
    R. Halim, M. K. Danquah and P. A. Webley, Biotechnol. Adv., 2012, 30, (3), 709 LINK https://doi.org/10.1016/j.biotechadv.2012.01.001
  3. 3.
    R. Subramaniam, S. Dufreche, M. Zappi and R. Bajpai, J. Ind. Microbiol. Biotechnol., 2010, 37, (12), 1271 LINK https://doi.org/10.1007/s10295-010-0884-5
  4. 4.
    L. Brennan and P. Owende, Renew. Sustain. Energy Rev., 2010, 14, (2), 557 LINK https://doi.org/10.1016/j.rser.2009.10.009
  5. 5.
    E. Tapia V, A. Anschau, A. L. V. Coradini, T. T Franco and A. Deckmann, AMB Express, 2012, 2, 64 LINK https://doi.org/10.1186/2191-0855-2-64
  6. 6.
    M. Thiru, S. Sankh and V. Rangaswamy, Bioresour. Technol., 2011, 102, (22), 10436 LINK https://doi.org/10.1016/j.biortech.2011.08.102
  7. 7.
    S. Qadeer, A. Khalid, S. Mahmood, M. Anjum and Z. Ahmad, J. Clean. Prod., 2017, 168, 917 LINK https://doi.org/10.1016/j.jclepro.2017.09.093
  8. 8.
    A. Patel, D. Karageorgou, E. Rova, P. Katapodis, U. Rova, P. Christakopoulos and L. Matsakas, Microorganisms, 2020, 8, (3), 434 LINK https://doi.org/10.3390/microorganisms8030434
  9. 9.
    I. R. Sitepu, L. A. Garay, R. Sestric, D. Levin, D. E. Block, J. B. German and K. L. Boundy-Mills, Biotechnol. Adv., 2014, 32, (7), 1336 LINK https://doi.org/10.1016/j.biotechadv.2014.08.003
  10. 10.
    V. Sànchez i Nogué, B. A. Black, J. S. Kruger, C. A. Singer, K. J. Ramirez, M. L. Reed, N. S. Cleveland, E. R. Singer, X. Yi, R. Y. Yeap, J. G. Linger and G. T. Beckham, Green Chem., 2018, 20, (18), 4349 LINK https://doi.org/10.1039/c8gc01905c
  11. 11.
    M.-H. Liang and J.-G. Jiang, Prog. Lipid Res., 2013, 52, (4), 395 LINK https://doi.org/10.1016/j.plipres.2013.05.002
  12. 12.
    L. R. Kumar, R. Kaur, S. K. Yellapu, X. Zhang, and R. D. Tyagi, ‘Biodiesel Production From Oleaginous Microorganisms With Wastes as Raw Materials’, in “Biofuels: Alternative Feedstocks and Conversion Processes for the Production of Liquid and Gaseous Biofuels”, 2nd Edn., eds. A. Pandey, C. Larroche, C.-G. Dussap, E. Gnansounou, S. K. Khanal and S. Ricke, ch. 32, Elsevier Inc, San Diego, USA, 2019, pp. 661–690 LINK https://doi.org/10.1016/B978-0-12-816856-1.00027-0
  13. 13.
    C. Zhao, T. Brück and J. A. Lercher, Green Chem., 2013, 15, (7), 1720 LINK https://doi.org/10.1039/c3gc40558c
  14. 14.
    T. Marker, J. Petri, T. Kalnes, M. McCall, D. Mackowiak, B. Jerosky, B. Reagan, L. Nemeth, M. Krawczyk, S. Czerni, D. Elliott and D. Shonnard, “Opportunities for Biorenewables in Oil Refineries: Final Technical Report”, DOE Award No. DE-FG36-05GO15085, UOP LLC, Des Plaines, USA, 2005, 60 pp LINK https://doi.org/10.2172/861458
  15. 15.
    Y. Liu, R. Sotelo-Boyás, K. Murata, T. Minowa and K. Sakanishi, Energy Fuels, 2011, 25, (10), 4675 LINK https://doi.org/10.1021/ef200889e
  16. 16.
    M. C. Vásquez, E. E. Silva and E. F. Castillo, Biomass Bioenergy, 2017, 105, 197 LINK https://doi.org/10.1016/j.biombioe.2017.07.008
  17. 17.
    R. Sotelo-Boyás, F. Trejo-Zárraga, and F. de Jesús Hernández-Loyo, ‘Hydroconversion of Triglycerides into Green Liquid Fuels’, in “Hydrogenation”, ed. I. Karamé, ch. 8, InTechOpen Ltd, London, UK, 2012, pp 187–216 LINK https://doi.org/10.5772/48710
  18. 18.
    P. Šimáček, D. Kubička, I. Kubičková, F. Homola, M. Pospíšil and J. Chudoba, Fuel, 2011, 90, (7), 2473 LINK https://doi.org/10.1016/j.fuel.2011.03.013
  19. 19.
    I. Kubičková, M. Snåre, K. Eränen, P. Mäki-Arvela and D. Yu. Murzin, Catal. Today, 2005, 106, (1–4), 197 LINK https://doi.org/10.1016/j.cattod.2005.07.188
  20. 20.
    I. Kubičková and D. Kubička, Waste Biomass Valoriz., 2010, 1, (3), 293 LINK https://doi.org/10.1007/s12649-010-9032-8
  21. 21.
    D. Kubička and J. Horáček, Appl. Catal. A: Gen., 2011, 394, (1–2), 9 LINK https://doi.org/10.1016/j.apcata.2010.10.034
  22. 22.
    M. Toba, Y. Abe, H. Kuramochi, M. Osako, T. Mochizuki and Y. Yoshimura, Catal. Today, 2011, 164, (1), 533 LINK https://doi.org/10.1016/j.cattod.2010.11.049
  23. 23.
    C. Wang, Z. Tian, L. Wang, R. Xu, Q. Liu, W. Qu, H. Ma and B. Wang, ChemSusChem, 2012, 5, (10), 1974 LINK https://doi.org/10.1002/cssc.201200219
  24. 24.
    C. Wang, Q. Liu, J. Song, W. Li, P. Li, R. Xu, H. Ma and Z. Tian, Catal. Today, 2014, 234, 153 LINK https://doi.org/10.1016/j.cattod.2014.02.011
  25. 25.
    M. Rabaev, M. V. Landau, R. Vidruk-Nehemya, A. Goldbourt and M. Herskowitz, J. Catal., 2015, 332, 164 LINK https://doi.org/10.1016/j.jcat.2015.10.005
  26. 26.
    M. Rabaev, M. V Landau, R. Vidruk-Nehemya, V. Koukouliev, R. Zarchin and M. Herskowitz, Fuel, 2015, 161, 287 LINK https://doi.org/10.1016/j.fuel.2015.08.063
  27. 27.
    M. Mohammad, T. K. Hari, Z. Yaakob, Y. C. Sharma and K. Sopian, Renew. Sustain. Energy Rev., 2013, 22, 121 LINK https://doi.org/10.1016/j.rser.2013.01.026
  28. 28.
    C. Yang, R. Li, C. Cui, S. Liu, Q. Qiu, Y. Ding, Y. Wu and B. Zhang, Green Chem., 2016, 18, (13), 3684 LINK https://doi.org/10.1039/c6gc01239f
  29. 29.
    M.-Y. Choo, L. E. Oi, P. L. Show, J.-S. Chang, T. C. Ling, E.-P. Ng, S. M. Phang and J. C. Juan, J. Taiwan Inst. Chem. Eng., 2017, 79, 116 LINK https://doi.org/10.1016/j.jtice.2017.06.028
  30. 30.
    X. Li, X. Luo, Y. Jin, J. Li, H. Zhang, A. Zhang and J. Xie, Renew. Sustain. Energy Rev., 2018, 82, (3), 3762 LINK https://doi.org/10.1016/j.rser.2017.10.091
  31. 31.
    T. Dong, E. P. Knoshaug, P. T. Pienkos and L. M. L. Laurens, Appl. Energy, 2016, 177, 879 LINK https://doi.org/10.1016/j.apenergy.2016.06.002
  32. 32.
    J. S. Kruger, N. S. Cleveland, R. Y. Yeap, T. Dong, K. J. Ramirez, N. J. Nagle, A. C. Lowell, G. T. Beckham, J. D. McMillan and M. J. Biddy, ACS Sustain. Chem. Eng., 2018, 6, (3), 2921 LINK https://doi.org/10.1021/acssuschemeng.7b01874
  33. 33.
    T. Dong, S. Van Wychen, N. Nagle, P. T. Pienkos and L. M. L. Laurens, Algal Res., 2016, 18, 69 LINK https://doi.org/10.1016/j.algal.2016.06.004
  34. 34.
    T. Dong, Q. Fei, M. Genelot, H. Smith, L. M. L. Laurens, M. J. Watson and P. T. Pienkos, Energy Convers. Manag., 2017, 140, 62 LINK https://doi.org/10.1016/j.enconman.2017.02.075
  35. 35.
    E. Ryckebosch, K. Muylaert and I. Foubert, J. Am. Oil Chem. Soc., 2012, 89, (2), 189 LINK https://doi.org/10.1007/s11746-011-1903-z
  36. 36.
    E. Ryckebosch, C. Bruneel, R. Termote-Verhalle, K. Muylaert and I. Foubert, Algal Res., 2014, 3, 36 LINK https://doi.org/10.1016/j.algal.2013.11.001
  37. 37.
    Y. Li, F. G. Naghdi, S. Garg, T. C. Adarme-Vega, K. J. Thurecht, W. A. Ghafor, S. Tannock and P. M. Schenk, Microb. Cell Fact., 2014, 13, 14 LINK https://doi.org/10.1186/1475-2859-13-14
  38. 38.
    X. Ren, X. Zhao, F. Turcotte, J.-S. Deschênes, R. Tremblay and M. Jolicoeur, Microb. Cell Fact., 2017, 16, 26 LINK https://doi.org/10.1186/s12934-017-0633-9
  39. 39.
    L. M. Wendt, B. D. Wahlen, E. P. Knoshaug, N. J. Nagle, T. Dong, R. Spiller, B. Panczak, S. Van Wychen, T. A. Dempster, H. Gerken and P. T. Pienkos, ACS Sustain. Chem. Eng., 2020, 8, (35), 13310 LINK https://doi.org/10.1021/acssuschemeng.0c03790
  40. 40.
    E. Ryckebosch, S. P. C. Bermúdez, R. Termote-Verhalle, C. Bruneel, K. Muylaert, R. Parra-Saldivar and I. Foubert, J. Appl. Phycol., 2013, 26, (3), 1501 LINK https://doi.org/10.1007/s10811-013-0189-y
  41. 41.
    F. Thevenieau and J.-M. Nicaud, OCL, 2013, 20, (6), D603 LINK https://doi.org/10.1051/ocl/2013034
  42. 42.
    G. F. Ferreira, L. F. Ríos Pinto, R. M. Filho and L. V Fregolente, Renew. Sustain. Energy Rev., 2019, 109, 448 LINK https://doi.org/10.1016/j.rser.2019.04.052
  43. 43.
    E. M. Fakhry and D. M. El Maghraby, J. Water Resour. Prot., 2013, 05, (09), 894 LINK https://doi.org/10.4236/jwarp.2013.59091
  44. 44.
    R. A. Ahmed, M. He, R. A. Aftab, S. Zheng, M. Nagi, R. Bakri and C. Wang, Sci. Rep., 2017, 7, 8118 LINK https://doi.org/10.1038/s41598-017-07540-x
  45. 45.
    X. Liu, J. Sheng and R. Curtiss, Proc. Natl. Acad. Sci., 2011, 108, (17), 6899 LINK https://doi.org/10.1073/pnas.1103014108
  46. 46.
    E. P. Knoshaug, S. Van Wychen, A. Singh and M. Zhang, Biofuel Res. J., 2018, 5, (2), 800 LINK https://doi.org/10.18331/brj2018.5.2.3
  47. 47.
    S. Santala, E. Efimova, V. Kivinen, A. Larjo, T. Aho, M. Karp and V. Santala, Microb. Cell Fact., 2011, 10, 36 LINK https://doi.org/10.1186/1475-2859-10-36
  48. 48.
    R. Kalscheuer, T. Stöveken, U. Malkus, R. Reichelt, P. N. Golyshin, J. S. Sabirova, M. Ferrer, K. N. Timmis and A. Steinbüchel, J. Bacteriol., 2007, 189, (3), 918 LINK https://doi.org/10.1128/jb.01292-06
  49. 49.
    I. Voss and A. Steinbüchel, Appl. Microbiol. Biotechnol., 2001, 55, (5), 547 LINK https://doi.org/10.1007/s002530000576
  50. 50.
    A. Arabolaza, E. Rodriguez, S. Altabe, H. Alvarez and H. Gramajo, Appl. Environ. Microbiol., 2008, 74, (9), 2573 LINK https://doi.org/10.1128/aem.02638-07
  51. 51.
    G. Venkata Subhash and S. Venkata Mohan, Bioresour. Technol., 2011, 102, (19), 9286 LINK https://doi.org/10.1016/j.biortech.2011.06.084
  52. 52.
    H. Gema, A. Kavadia, D. Dimou, V. Tsagou, M. Komaitis and G. Aggelis, Appl. Microbiol. Biotechnol., 2002, 58, (3), 303 LINK https://doi.org/10.1007/s00253-001-0910-7
  53. 53.
    S. U. Ahmed, S. K. Singh, A. Pandey, S. Kanjilal and R. B. N. Prasad, Food Technol. Biotechnol., 2006, 44, (2), 283 LINK https://hrcak.srce.hr/109905
  54. 54.
    A.-N. Vamvakaki, I. Kandarakis, S. Kaminarides, M. Komaitis and S. Papanikolaou, Eng. Life Sci., 2010, 10, (4), 348 LINK https://doi.org/10.1002/elsc.201000063
  55. 55.
    J. Hussain, Z. Ruan, I. A. Nascimento, Y. Liu and W. Liao, Bioresour. Technol., 2014, 169, 768 LINK https://doi.org/10.1016/j.biortech.2014.06.074
  56. 56.
    E. Carota, S. Crognale, A. D’Annibale and M. Petruccioli, Process Saf. Environ. Prot., 2018, 117, 143 LINK https://doi.org/10.1016/j.psep.2018.04.022
  57. 57.
    R. F. Wilson, ‘Seed Composition’, in “Soybeans: Improvement, Production, and Uses”, eds. H. R. Boerma and J. E. Specht, 3rd Edn., ch. 13, Vol. 16, American Society of Agronomy, Inc, Crop Science Society of America, Inc, Soil Science Society of America Inc, Madison, USA, 2004, pp. 621–677 LINK https://doi.org/10.2134/agronmonogr16.3ed.c13
  58. 58.
    T. E. Clemente and E. B. Cahoon, Plant Physiol., 2009, 151, (3), 1030 LINK https://doi.org/10.1104/pp.109.146282
  59. 59.
    D. Barrera-Arellano, A. P. Badan-Ribeiro and S. O. Serna-Saldivar, ‘Corn Oil: Composition, Processing, and Utilization’, in “Corn: Chemistry and Technology”, 3rd Edn., ch. 21, Elsevier Inc, Cambridge, USA, 2019, pp. 593–613 LINK https://doi.org/10.1016/B978-0-12-811971-6.00021-8
  60. 60.
    L. Chen, T. Liu, W. Zhang, X. Chen and J. Wang, Bioresour. Technol., 2012, 111, 208 LINK https://doi.org/10.1016/j.biortech.2012.02.033
  61. 61.
    L. Yao, J. A. Gerde and T. Wang, J. Am. Oil Chem. Soc., 2012, 89, (12), 2279 LINK https://doi.org/10.1007/s11746-012-2124-9
  62. 62.
    E. H. Pryde, ‘Composition of Soybean Oil’, in “Handbook of Soy Oil Processing and Utilization”, eds. D. R. Erikson, E. H. Pryde, O. L. Brekke, T. L. Mounts and R. A. Falb, Ch. 2, American Soybean Assoc, St. Louis, USA and American Oil Chemists Soc, Champaign, USA, 1980, pp. 13-31
  63. 63.
    T. Dong, D. Gao, C. Miao, X. Yu, C. Degan, M. Garcia-Pérez, B. Rasco, S. S. Sablani and S. Chen, Energy Convers. Manag., 2015, 105, 1389 LINK https://doi.org/10.1016/j.enconman.2015.06.072
  64. 64.
    A. de Jesus, A. V Zmozinski, J. A. Barbará, M. G. R. Vale and M. M. Silva, Energy Fuels, 2010, 24, (3), 2109 LINK https://doi.org/10.1021/ef9014235
  65. 65.
    W. Zschau, Eur. J. Lipid Sci. Technol., 2001, 103, (8), 505 LINK https://doi.org/10.1002/1438-9312(200108)103:8<505::aid-ejlt505>3.0.co;2-7
  66. 66.
    F. Hussin, M. K. Aroua and W. M. A. W. Daud, Chem. Eng. J., 2011, 170, (1), 90 LINK https://doi.org/10.1016/j.cej.2011.03.065
  67. 67.
    A. A. Aachary, J. Liang, A. Hydamaka, N. A. M. Eskin and U. Thiyam-Holländer, LWT - Food Sci. Technol., 2016, 72, 439 LINK https://doi.org/10.1016/j.lwt.2016.05.011
  68. 68.
    D. D. Brooks, S. A. Brophy and G. R. Goss, Oil-Dri Corporation of America, ‘Oil Bleaching Method and Composition for Same’, US Patent 1991; 5,004,570
  69. 69.
    V. Kuuluvainen, P. Mäki-Arvela, A.-R. Rautio, K. Kordas, J. Roine, A. Aho, B. Toukoniitty, H. Österholm, M. Toivakka and D. Y. Murzin, J. Chem. Technol. Biotechnol., 2015, 90, (9), 1579 LINK https://doi.org/10.1002/jctb.4621
  70. 70.
    L. L. Diosady, Int. J. Appl. Sci. Eng., 2005, 3, (2), 81
  71. 71.
    E. Santillan-Jimenez, R. Pace, S. Marques, T. Morgan, C. McKelphin, J. Mobley and M. Crocker, Fuel, 2016, 180, 668 LINK https://doi.org/10.1016/j.fuel.2016.04.079
  72. 72.
    J. S. Kruger, E. D. Christensen, T. Dong, S. Van Wychen, G. M. Fioroni, P. T. Pienkos and R. L. McCormick, Energy Fuels, 2017, 31, (10), 10946 LINK https://doi.org/10.1021/acs.energyfuels.7b01867
  73. 73.
    A. Sathish and R. C. Sims, Bioresour. Technol., 2012, 118, 643 LINK https://doi.org/10.1016/j.biortech.2012.05.118
  74. 74.
    T. Li, J. Xu, H. Wu, G. Wang, S. Dai, J. Fan, H. He and W. Xiang, Marine Drugs, 2016, 14, (9), 162 LINK https://doi.org/10.3390/md14090162
  75. 75.
    S. Paisan, P. Chetpattananondh and S. Chongkhong, J. Environ. Chem. Eng., 2017, 5, (5), 5115 LINK https://doi.org/10.1016/j.jece.2017.09.045
  76. 76.
    Y. C. Sharma, M. Yadav and S. N. Upadhyay, Biofuels, Bioprod. Biorefining, 2019, 13, (1), 174 LINK https://doi.org/10.1002/bbb.1937
  77. 77.
    O. Zufarov, Š. Schmidt and S. Sekretár, Acta Chim. Slovaca, 2008, 1, 321–328
  78. 78.
    S. Franklin, A. Somanchi, K. Espina, G. Rudenko and P. Chua, Solazyme Inc, Renewable Fuels Produced from Oleaginous Microorganisms, US Patent 9,062,294; 2015
  79. 79.
    A. Kale, L. C. Zullo and S. Shinde, Heliae Development LLC, Methods of and Systems for Producing Biofuels from Algal Oil, US Patent 8,313,648; 2012
  80. 80.
    A. Kale, Extraction of Polar Lipids by a Two Solvent Method, US Patent 8,475,660; 2013
  81. 81.
    H. L. Barnebey and A. C. Brown, J. Am. Oil Chem. Soc., 1948, 25, (3), 95 LINK https://doi.org/10.1007/bf02579733
  82. 82.
    E. N. Coppola, S. Nana and C. Red, Applied Research Associates, Inc, ‘Hydrothermal Cleanup Process’, US Patent 10,071,322; 2018
  83. 83.
    A. Lawal, J. Manganaro, B. Goodall and R. Farrauto, “Pt-Based Bi-Metallic Monolith Catalysts for Partial Upgrading of Microalgae Oil”, US DOE Office of Energy Efficiency and Renewable Energy (EERE), Washington, DC, USA, 2015, 130 pp LINK https://doi.org/10.2172/1344891
  84. 84.
    I. Hachemi, K. Jeništová, P. Mäki-Arvela, N. Kumar, K. Eränen, J. Hemming and D. Y. Murzin, Catal. Sci. Technol., 2016, 6, (5), 1476 LINK https://doi.org/10.1039/c5cy01294e
  85. 85.
    I. Hachemi, N. Kumar, P. Mäki-Arvela, J. Roine, M. Peurla, J. Hemming, J. Salonen and D. Y. Murzin, J. Catal., 2017, 347, 205 LINK https://doi.org/10.1016/j.jcat.2016.12.009
  86. 86.
    C. V. Viêgas, I. Hachemi, S. P. Freitas, P. Mäki-Arvela, A. Aho, J. Hemming, A. Smeds, I. Heinmaa, F. B. Fontes, D. C. da Silva Pereira, N. Kumar, D. A. G. Aranda and D. Y. Murzin, Fuel, 2015, 155, 144 LINK https://doi.org/10.1016/j.fuel.2015.03.064
  87. 87.
    L. Zhou and A. Lawal, Energy Fuels, 2015, 29, (1), 262 LINK https://doi.org/10.1021/ef502258q
  88. 88.
    L. Zhou and A. Lawal, Catal. Sci. Technol., 2016, 6, (5), 1442 LINK https://doi.org/10.1039/c5cy01307k
  89. 89.
    R. Loe, E. Santillan-Jimenez, T. Morgan, L. Sewell, Y. Ji, S. Jones, M. A. Isaacs, A. F. Lee and M. Crocker, Appl. Catal. B: Environ., 2016, 191, 147 LINK https://doi.org/10.1016/j.apcatb.2016.03.025
  90. 90.
    E. Santillan-Jimenez, T. Morgan, R. Loe and M. Crocker, Catal. Today, 2015, 258, (2), 284 LINK https://doi.org/10.1016/j.cattod.2014.12.004
  91. 91.
    M. H. Wilson, J. Groppo, A. Placido, S. Graham, S. A. Morton, E. Santillan-Jimenez, A. Shea, M. Crocker, C. Crofcheck and R. Andrews, Appl. Petrochem. Res., 2014, 4, (1), 41 LINK https://doi.org/10.1007/s13203-014-0052-3
  92. 92.
    B. Peng, Y. Yao, C. Zhao and J. A. Lercher, Angew. Chem. Int. Ed., 2012, 51, (9), 2072 LINK https://doi.org/10.1002/anie.201106243
  93. 93.
    B. Peng, X. Yuan, C. Zhao and J. A. Lercher, J. Am. Chem. Soc., 2012, 134, (22), 9400 LINK https://doi.org/10.1021/ja302436q
  94. 94.
    W. Song, C. Zhao and J. A. Lercher, Chem. Eur. J., 2013, 19, (30), 9833 LINK https://doi.org/10.1002/chem.201301005
  95. 95.
    H. S. H. Nguyen, P. Mäki-Arvela, U. Akhmetzyanova, Z. Tišler, I. Hachemi, A. Rudnäs, A. Smeds, K. Eränen, A. Aho, N. Kumar, J. Hemming, M. Peurla and D. Y. Murzin, J. Chem. Technol. Biotechnol., 2017, 92, (4), 741 LINK https://doi.org/10.1002/jctb.5158
  96. 96.
    H. S. H. Nguyen, I. Hachemi, A. Rudnas, P. Mäki-Arvela, A. Smeds, A. Aho, J. Hemming, M. Peurla and D. Y. Murzin, Ind. Eng. Chem. Res., 2016, 55, (40), 10626 LINK https://doi.org/10.1021/acs.iecr.6b03041
  97. 97.
    E. P. Knoshaug, A. Mohagheghi, N. J. Nagle, J. J. Stickel, T. Dong, E. M. Karp, J. S. Kruger, D. G. Brandner, L. P. Manker, N. A. Rorrer, D. A. Hyman, E. D. Christensen and P. T. Pienkos, Green Chem., 2018, 20, (2), 457 LINK https://doi.org/10.1039/c7gc02295f
  98. 98.
    H. J. Robota, J. C. Alger and L. Shafer, Energy Fuels, 2013, 27, (2), 985 LINK https://doi.org/10.1021/ef301977b
  99. 99.
    V. K. Soni, P. R. Sharma, G. Choudhary, S. Pandey and R. K. Sharma, ACS Sustain. Chem. Eng., 2017, 5, (6), 5351 LINK https://doi.org/10.1021/acssuschemeng.7b00659
  100. 100.
    J. Fu, C. Yang, J. Wu, J. Zhuang, Z. Hou and X. Lu, Fuel, 2015, 139, 678 LINK https://doi.org/10.1016/j.fuel.2014.09.025
  101. 101.
    K. Kandel, J. W. Anderegg, N. C. Nelson, U. Chaudhary and I. I. Slowing, J. Catal., 2014, 314, 142 LINK https://doi.org/10.1016/j.jcat.2014.04.009
  102. 102.
    S. Tang, Z. Shi, X. Tang and X. Yang, Green Chem., 2019, 21, (12), 3413 LINK https://doi.org/10.1039/c9gc00673g
  103. 103.
    M. K. Poddar, M. Anand, S. A. Farooqui, G. J. O. Martin, M. R. Maurya and A. K. Sinha, Biomass Bioenergy, 2018, 119, 31 LINK https://doi.org/10.1016/j.biombioe.2018.08.011
  104. 104.
    W.-C. Wang, E. Allen, A. A. Campos, R. K. Cade, L. Dean, M. Dvora, J. G. Immer, S. Mixson, S. Srirangan, M.-L. Sauer, S. Schreck, K. Sun, N. Thapaliya, C. Wilson, J. Burkholder, A. M. Grunden, H. H. Lamb, H. Sederoff, L. F. Stikeleather and W. L. Roberts, Environ. Prog. Sustain. Energy, 2013, 32, (4), 916 LINK https://doi.org/10.1002/ep.11855
  105. 105.
    T. C. Schulz, M. Oelschlager, S. T. Thompson, W. F. J. Vermaas, D. R. Nielsen and H. H. Lamb, Sustain. Energy Fuels, 2018, 2, (4), 882 LINK https://doi.org/10.1039/c7se00558j
  106. 106.
    T. J. Czartoski, R. Perkins, J. L. Villanueva and G. Richards, Kohn & Assoc PLLC, Algae Biomass Fractionation, US Patent Appl. 2010/233,761
  107. 107.
    J. L. Wagner, V. P. Ting and C. J. Chuck, Fuel, 2014, 130, 315 LINK https://doi.org/10.1016/j.fuel.2014.04.048
  108. 108.
    R. W. Jenkins, L. A. Sargeant, F. M. Whiffin, F. Santomauro, D. Kaloudis, P. Mozzanega, C. D. Bannister, S. Baena and C. J. Chuck, ACS Sustain. Chem. Eng., 2015, 3, (7), 1526 LINK https://doi.org/10.1021/acssuschemeng.5b00228
  109. 109.
    M. Bystrzanowska, P. Petkov and M. Tobiszewski, ACS Sustain. Chem. Eng., 2019, 7, (22), 18434 LINK https://doi.org/10.1021/acssuschemeng.9b04230
  110. 110.
    M. Marafi and E. Furimsky, Energy Fuels, 2017, 31, (6), 5711 LINK https://doi.org/10.1021/acs.energyfuels.7b00471
  111. 111.
    B. Zhang, J. Wu, C. Yang, Q. Qiu, Q. Yan, R. Li, B. Wang, J. Wu and Y. Ding, BioEnergy Res., 2018, 11, (3), 689 LINK https://doi.org/10.1007/s12155-018-9927-y
  112. 112.
    B. Donnis, R. G. Egeberg, P. Blom and K. G. Knudsen, Top. Catal., 2009, 52, (3), 229 LINK https://doi.org/10.1007/s11244-008-9159-z

Acknowledgements

This work was by the National Renewable Energy Laboratory and financially supported by the US Department of Energy under Contract No. DE-AC36-08GO28308 with the National Renewable Energy Laboratory, as part of the DOE Office of Energy Efficiency and Renewable Energy, Bioenergy Technologies Office. The views expressed in the article do not necessarily represent the views of the DOE or the US Government. The US Government retains and the publisher, by accepting the article for publication, acknowledges that the US Government retains a nonexclusive, paid-up, irrevocable, world-wide license to publish or reproduce the published form of this work, or allow others to do so, for US Government purposes.

The Authors


Jacob S. Kruger is a research engineer at NREL. Prior to joining NREL in 2013. He studied biodiesel chemistry at Hamline University, Saint Paul, Minnesota, USA (BA, 2007), autothermal reforming of renewable feedstocks at the University of Minnesota, USA (PhD, 2011) and carbohydrate dehydration over zeolite catalysts at the University of Delaware, USA (postdoctoral research, 2013). His current work involves catalytic processing of lignocellulose and other renewable feedstocks, with a focus on conversion of lignin residues and microbial biomass to high-value products.


Eric P. Knoshaug is a Group Manager/Scientist at NREL with a BA from the University of Colorado Boulder, USA, in Environmental Studies/Economics and an MS in Microbiology from Oregon State University, Corvallis, Oregon, USA. His current work involves biological conversion of pretreated algal biomass to fuel and chemical precursors, genetic engineering of green microalgae and data analytics of pilot-scale algae cultivation.


Tao Dong is a research engineer at NREL. Prior to joining NREL in 2013, Dong studied Food Chemistry at Jiangnan University, China, (BA, 2005; MS, 2008) and Biological Systems Engineering at Washington State University, USA (PhD, 2013). His current work involves downstream processing of algal biomass to produce precursors for fuel and value-added coproducts, and development of fully renewable bio-based polymers for industrial applications.


Tobias C. Hull is a PhD candidate in the Advanced Energy Systems programme at the Colorado School of Mines, USA. Previously, he studied chemical engineering at the Rochester Institute of Technology, New York, USA (BS, 2018). His thesis aims to explore the role of bioenergy in the ongoing energy transition.


Philip T. Pienkos earned a BS in Honours Biology at the University of Illinois, USA, and his PhD in Molecular Biology at the University of Wisconsin–Madison, USA. He co-founded two companies (Molecular Logix and Celgene), prior to joining NREL in 2007 as a Principal Group Manager. At NREL, Pienkos worked in various aspects of cellulosic ethanol and algal biofuels production, and helped establish new biofuels research and development areas including biological methane upgrading and waste to energy. He is currently the CEO of Polaris Renewables, established as a platform for his consulting business, and Emeritus Scientist at NREL.

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