*2.3. Moisture Analysis*

Moisture was determined in triplicate in the oven (SP 200, SP Labor®, São Paulo, Brazil), at 65 ± 1 ◦C, for approximately 72 h [29].

#### *2.4. Extraction of Carotenoids*

The carotenoid pigments were extracted from the lyophilized material (peel + pulp), according to the recommended procedures by Rodrigues-Amaya et al. [30]. Three grams of the edible portion of the samples (peel + pulp) were crushed by the use of a mortar and pestle, and a few drops of distilled water were added and extracted to color exhaustion with 20 mL of acetone (7 times) in an ultrasonic bath (Labsonic LBS 1-H22.5, Treviglio, Bergamo, Italy) for 10 min each time. Then, the extracts were individually centrifuged (Awel MF20-R, Multifunction Refrigerated Centrifuge, Blain, France) at 4000 rpm, 5 ◦C, for 10 min in order to withdraw clear solution on the top. The acetonic extracts were pooled together was concentrated to about 25 mL, in a rotary evaporator (Buchi-heating bath B-491, Buchi, Milan, Italy) at temperature below 35 ◦C. The dry product was diluted with equal volumes (25 mL) of a mixture of ethyl ether and hexane (1:1) and distilled water (50 mL) and worked up with a separating funnel. The lipofilic phase, cleared by the hydrophilic impurities, was evaporated to dryness using a rotary evaporator (Buchi-heating bath B-491 at 35 ◦C, and the residue was dissolved in 2 mL of MeOH/MTBE (1:1) and filtered in filter units (PTFE, 0.45μm, 13mm, Sigma Aldrich, Milan, Italy) prior to HPLC analysis. Samples were stored at −20 ◦C until they were analyzed.

#### *2.5. Analysis of Carotenoids by HPLC-DAD-APCI-MS*

The analysis was performed on an HPLC system (Shimadzu, Kyoto, Japan) equipped with a CBM-20A controller, two LC-20AD pumps, a DGU-20A3R deaerator, a SIL-20AC autosampler, a CTO 20AC column oven and an SPD-M20A photo diode array detector. The data were processed with the Labsolution software. For MS analysis a mass spectrometer detector (LCMS-8040) was used, equipped with an APCI (atmospheric pressure chemical ionization) interface, both in positive and negative ionization mode.

The column used was YMC C30 (250 mm × 4.6 mm × 5 μm); the mobile phases: MeOH/MTBE/H2O (81:15:4, solvent A), MeOH/MTBE/H2O (6:90:4, solvent B); the linear gradient used was: 0–100% B from 0 to 140 min. The column temperature was maintained at 30 ◦C. The flow was 0.8 mL/min and the injection volume was 20 μL.

The UV-Vis spectra were acquired in the range 220–700 nm, while the chromatograms were extracted at 450 nm (sampling frequency: 4.16 Hz, time constant: 0.64 s). The MS was set up as follows: Scan, both APCI positive (+) and negative (-); atomized gas flow (N2): 2.0 L/min; drying gas flow: 5 L/min; Time of the event: 0.06 s; range *m*/*z*: 300–1200; interface temperature: 350 ◦C; Desolvation line (DL) temperature: 300 ◦C; thermal block: 300 ◦C. The samples were analyzed in triplicate.

#### *2.6. Identification and Quantification of Carotenoids*

Carotenoids were identified by their UV-Vis spectra, MS spectra, elution order, comparison with the available standard and literature data.

The kumquat carotenoids quantification was performed from the analytical curves. External standards quantitative determination of each compound was performed using all reference materials listed in Section 2.1, in the concentration range from 5 to 50 μg/mL at six concentration levels. The results were obtained from the average of three determinations and the CV% was below 8% in all the LC measurements. The *R* coefficient for the calibration curves was always above 0.9962, with LOD and LOQ values of 0.07 and 0.22 ppm for β-carotene, 0.1 and 0.33 ppm for β-cryptoxanthin, 0.06 and 0.18 ppm for lutein, 0.08 and 0.3 ppm for zeaxanthin, and 0.12 and 0.24 ppm for physalein, respectively.

#### **3. Results and Discussion**

#### *3.1. Carotenoids Qualitative Profile of Brazilian Kumquat*

Figure 2 shows the chromatographic profile of the carotenoid composition in not saponified kumquat fruits extracts. The identified compounds are shown in Table 1, together with the UV–Vis and MS spectra information. In Figure 3 are reported the UV-Vis (PDA) and mass spectrum of β-citraurin-laurate and β-citraurin-myristate, detected in the kumquat carotenoid extracts. It can be appreciated that the esterification does not affect the PDA spectra of β-citraurin.

**Figure 2.** Chromatographic profile of native carotenoids of kumquat from Brazil: peel + pulp. Identification of the compounds are in Table 1.

**Table 1.** Compounds identification for Figure 2 (kumquat from Brazil: peel + pulp).

Rt: retention time; PDA: photodiode array; λnm: wavelength of maximum absorption; MS: mass spectrometry; APCI: atmospheric pressure chemical ionization.

**Figure 3.** UV-Vis (PDA) and mass spectrum (APCI negative) of β-citraurin-laurate (**A**) and β-citraurinmyristate (**B**) of kumquat from Brazil.

Moreover, both the molecular ions [M]−• at respectively *m*/*z* 614 and *m*/*z* 642 relative to the β-citraurin-laurate and β-citraurin-myristate esters, obtained in the negative APCI mode, are also clearly shown in the same figure. Eleven different carotenoids were identified in kumquat from Brazil; four different β-citraurin esters and three β-cryptoxanthin esters were identified.

The predominant carotenoids were β-citraurin-laurate and β-cryptoxanthin-laurate, both esterified with lauric acid. The chemical structures of these carotenoids are shown in Figure 4. Interestingly, no free β-citraurin was detected in the present study.

**Figure 4.** Chemical structures of β-cryptoxanthin-laurate and β-citraurin-laurate, main carotenoids detected in kumquat.

Shirra et al. [9] detected only 4 carotenoids in kumquat from Italy without saponification (β-carotene, β-cryptoxanthin, lutein and zeaxanthin), which were also detected in the present study. In contrast to our study, Agos et al. [18] reported only β-citraurin and β-cryptoxanthin in the free form in kumquat from Hungary. However, these researchers used saponification after the extraction process and did not quantify the carotenoid esters. Saponification may result in destruction or structural transformation of carotenoids [31]. Huyskens et al. [19] studied the qualitative composition of kumquat carotenoids from Israel by thin-layer chromatography and reported several carotenoids, including β-citraurin, β-cryptoxanthin, lutein, β-carotene and zeaxanthin which were, also determined in the present study. In addition, Huyskens et al. [19] reported that violaxanthin was the predominant component in kumquat from Israel, whereas in our study β-citraurin-laurate was the major component.

Esterification with saturated fatty acids improves the stability of xanthophylls such as β-citraurin and β-cryptoxanthin against heat and UV light, but does not affect their antioxidant activity [32]. During the storage and processing of the fruits the xanthophyll esters were more stable than the free xanthophyll [33]. The pigment β-citraurin is responsible for the citrus reddish color, it derives from of β-cryptoxanthin or zeaxanthin and accumulates in some citrus varieties [34,35]. In these fruits, the accumulation of β-citraurin is not a common event, it is observed only in the flavedos of some varieties during fruit ripening [34]. Frutita, a tropical fruit from Panama, showed a very high content of β-citraurin [36].

Recent studies have shown that β-cryptoxanthin and β-citraurin esterified with lauric acid, myristic acid and palmitic acid are found in the Mandarin Satsuma, [35].

The dietary intake of β-cryptoxanthin has been shown to prevent and reduce some pathologies such as: cancer, diabetes and rheumatism due to its antioxidant activity [37–39]. Breithaupt et al. [40] have verified that in chili, papaya, peach and persimmon, β-cryptoxanthin is mainly esterified with saturated fatty acids. Furthermore, the bioavailability of β-cryptoxanthin esters is comparable to the non-esterified form, since fatty acids can be effectively hydrolyzed by β-cryptoxanthin esters before intestinal absorption in the human body [40]. In several citrus varieties, the esterified form β-citraurin is found [35]. As already observed in another study [41], we have found β-cryptoxanthin and β-citraurin in both free and esterified forms.

#### *3.2. Carotenoids Quantitative Profile of Brazilian Kumquat*

In the present study, β-citraurin-laurate was the carotenoid found in the highest content in kumquat from Brazil (607.33 μg/100 g fresh matter, representing 27.80% of total carotenoids), followed by β-cryptoxanthin-laurate (552.59 μg/100 g fresh matter, representing 25.31% of total carotenoids). Thus, β-citraurin-laurate and β-cryptoxanthin-laurate represented 53.11% of the total carotenoids

found. The different forms of β-citraurin (4 components esterified with fatty acids) and β-cryptoxanthin (1 component in free form and 3 in esterified form) represented 84.34% of the carotenoids found in kumquat from Brazil. Forms of β-citraurin and β-cryptoxanthin esterified with fatty acids accounted for 79.54% of the total carotenoids in kumquat. These results show the importance of the study of the intact carotenoids composition in different matrices (Table 2).

**Table 2.** Carotenoid content in kumquat from Brazil (peel + pulp).

\* Mean of 3 repetitions ± standard deviation (SD). The carotenoid content in the fresh kumquat was calculated based on the average moisture content (*n* = 3) of the lyophilized fruit (14.08%) and the fresh fruit (81.16%).

β-carotene was found in smaller amounts (120.19 μg/100 g fresh matter), representing 5.50% of total carotenoids in kumquat, as well as free xanthophylls (β-cryptoxanthin, lutein and zeaxanthin), which represented 14.96% of total carotenoids (326.96 μg/100 g) (Table 2). β-carotene and β-cryptoxanthin possess provitamin A activity, playing a key role in human health [42]. Differently from our results, Wang et al. [20] reported that β-cryptoxanthin was the major carotenoid present in kumquat cultivated in Taiwan, while Schirra et al. [9] found lutein to be the major carotenoid present in kumquat from Italy.

Schirra et al. [9] found a lower concentration, in fresh matter, of β-carotene (33 μg/100 g), β-cryptoxanthin (26 μg/100 g), lutein (44 μg/100 g) and zeaxanthin (24 μg/100 g) in kumquat from Italy when compared to the concentrations of these carotenoids in kumquat from Brazil, reported here for the first time. Wang et al. [41] found contents in dry matter that were much lower than reported here, for lutein (9.9 μg/100 g), zeaxanthin (10.4 μg/100 g), β-cryptoxanthin (183 μg/100 g) and β-carotene (131 μg/100 g) in kumquat cultivated in Taiwan. Carotenoid composition and contents in fruits can be influenced by genetic factors, geographical regions, fruit processing, storage methods Giuffrida et al. [25] and environmental conditions Lu et al. [26], which can explain the differences found in these studies. Besides the climatic factors, the irrigation, soil conditions, fertilizers and herbicides may also affect the carotenoids accumulation [26]. Regarding the carotenoids content in food, Britton [43] have proposed the following classification ranges: low: 0–0.1 mg/100 g; moderate: 0.1–0.5 mg/100 g; high: 0.5-2 mg/100 g; very high: >2 mg/100 g. Thus, the total carotenoid content of kumquat from Brazil was very high (2185.16 μg/100 g).


**Yvan Torrente \*, Pamela Bella, Luana Tripodi, Chiara Villa and Andrea Farini \***

Stem Cell Laboratory, Department of Pathophysiology and Transplantation, University of Milan, Unit of Neurology, Fondazione IRCCS Cà Granda Ospedale Maggiore Policlinico, Dino Ferrari Center, 20122 Milan, Italy<EMAIL_ADDRESS>(P.B<EMAIL_ADDRESS>(L.T<EMAIL_ADDRESS>(C.V.)

**\*** Correspondence<EMAIL_ADDRESS>(Y.T<EMAIL_ADDRESS>(A.F.); Tel.: +39-0255033874 (Y.T.); +39-0255033852 (A.F.)

Received: 20 January 2020; Accepted: 11 February 2020; Published: 14 February 2020

**Abstract:** The insulin-like growth factor 2 receptor (IGF2R) plays a major role in binding and regulating the circulating and tissue levels of the mitogenic peptide insulin-like growth factor 2 (IGF2). IGF2/IGF2R interaction influences cell growth, survival, and migration in normal tissue development, and the deregulation of IGF2R expression has been associated with growth-related disease and cancer. IGF2R overexpression has been implicated in heart and muscle disease progression. Recent research findings suggest novel approaches to target IGF2R action. This review highlights recent advances in the understanding of the IGF2R structure and pathways related to muscle homeostasis.

**Keywords:** IGF2R; muscle homeostasis; inflammation; muscular dystrophy; pericytes


*2.1. Model with Distributed Parameters*

The number of newly infected individuals *J*(*t*) is determined by the rate of decrease of the number of susceptible individuals, *J*(*t*) = −*S* (*t*). Assuming that

$$N = S(t) + I(t) + R(t) + D(t) \tag{1}$$

is constant, where *I*(*t*) is the total number of infected at time *t* and *R*(*t*) and *D*(*t*) denote, respectively, recovered and dead, we can write

$$I(t) = \int\_0^t J(\eta)d\eta - R(t) - D(t). \tag{2}$$

Following conventional epidemiological models, we set

$$\frac{dS(t)}{dt} = -\beta \frac{S(t)}{N} \,\, I(t),$$

where *β* is the disease transmission rate. Let *r*(*η*) and *d*(*η*) be the recovery and death rates depending on the time-since-infection *η*. Then, the number of infected individuals who will recover at time *t* is given by the expression:

$$\frac{d\mathcal{R}(t)}{dt} = \int\_0^t r(t-\eta)f(\eta)d\eta.$$

and the number of infected individuals who will die at time *t*:

$$\frac{dD(t)}{dt} = \int\_0^t d(t-\eta)f(\eta)d\eta.$$

Differentiating Equality (1), we obtain

$$\frac{dI(t)}{dt} = \beta \frac{S(t)}{N} \left. I(t) - \int\_0^t r(t - \eta) I(\eta) d\eta - \int\_0^t d(t - \eta) I(\eta) d\eta. \right|$$

Thus, we obtain the following integro-differential equation model:

$$\frac{dS(t)}{dt}\_{\text{in}} = -\beta \frac{S(t)}{N} I(t) \quad (= -I(t)), \tag{3a}$$

$$\frac{dI(t)}{dt}\_{\text{min}} = \beta \frac{S(t)}{N} I(t) - \int\_0^t r(t-\eta)f(\eta)d\eta - \int\_0^t d(t-\eta)f(\eta)d\eta,\tag{3b}$$

$$\frac{d\mathcal{R}(t)}{dt} = -\int\_0^t r(t-\eta)f(\eta)d\eta,\tag{3c}$$

$$\frac{dD(t)}{dt} = -\int\_0^t d(t-\eta)f(\eta)d\eta,\tag{3d}$$

with the initial condition *S*(0) = *N*, *I*(0) = *I*0 > 0 (*I*0 is sufficiently small as compared to *N*), *R*(0) = 0, *D*(0) = 0.


**3.1.2 Batch processing**

#### <IP_ADDRESS> Preheating

Mechanically pressing crude soybean oil is preheated to 80 degrees Celsius after removal of impurities by filtration.

#### <IP_ADDRESS> Hydration

The amount of water added is determined by phospholipids content in the oil and the changes of phospholipids granules formed during heating and is normally 3.5 times (w/w) the content of phospholipids. The water added is usually boiling or 0.7% hot salt solution is used. The speed of adding water is determined by the water absorption velocity of phospholipids. The faster the latter the faster the former, and vice versa. When adding water, the stirring speed must be fast and is normally 80-100rpm at the beginning and is slowed down 20-30min later when large flocculent phospholipids granules are formed and the stirring is continued for another 20-30min. Then the liquid is left standing still to settle. The supernatant of the upper phase is dehydrated to produce refined oil while the oil foot of the lower phase need to be concentrated to obtain phospholipids products.

#### <IP_ADDRESS> Concentration

The hydrated phospholipids oil foot is drawn into the concentrating tank by vacuum and subjected to temperature rising and stirring. Vacuum dehydration of phospholipids occurs at about 80 degrees Celsius. When there is slight silk flash while stirring the fluidic phospholipids the moisture content is consistent with the specification. The moisture content is about 5%. Phospholipids after concentration is a brown semisolid and can be used in food, medicine and industry.

#### <IP_ADDRESS> Decoloration

Decoloration of concentrated phospholipids is needed for preparation of high quality phospholipids. The amount of 30% hydrogen peroxide added to the concentrating tank is 2%-2.5% (w/w) of the concentrated phospholipids. The phospholipids are decolored in the closed tank for 1h at 50 degrees Celsius without vacuum. Then turn on the vacuum pump and heat the mixture to 70 degrees Celsius. Dehydrate until there is no water in the water knock vessel. The decolored phospholipids are light brown.

Soybean Phospholipids 489

are fed into the double conic dryer with the amount of 1/3-1/2 of the whole dryer volume. The drying parameters are as follows: drying temperature 50-55 degrees Celsius, rotate speed 10rpm, vacuum -0.083--0.09 MPa, time 4-6h. Then light yellow powdery

The above method can be applied to prepare powdery phospholipids from such various raw materials as soybean, rapeseed, peanut and corn etc. as well as concentrated phospholipids prepared from hydrated oil foot and alkalized oil foot. The powdery phospholipids produced have a phospholipids content of 90%-98% due to the quality of the concentrated

<IP_ADDRESS> Preparation of powdery phospholipids of various purities from one kind of materials In acetone solvent extraction, the phospholipids purity increases with the increase of acetone amount and extraction times. The increase of phospholipids purity results in longer

If the purity of the powdery phospholipids obtained in <IP_ADDRESS> is 97%-98%, half of the phospholipids will be settled in 0.5-1h in the fourth extraction while the other half in 4h. The upper phospholipids solution of acetone is discharged when the time has passed 2.5-3h and centrifuged and dried. The purity of the phospholipids produced can reach up to over 99%. Acetone of 2 times the weight of concentrated phospholipids is added into the extraction tank with agitation. The extraction time is 0.5h and static settle time is 1.5-2h. The upper phospholipids solution of acetone is discharged and centrifuged and dried to obtain

The residual liquid is discharged, centrifuged and dried to produce phospholipids product

This method can produce phospholipids products with various purities due to the product purity obtained in <IP_ADDRESS> and discharging time to meet the market requirements, and make

Extraction temperature, pressure and time are important technological parameters of supercritical carbon dioxide extraction. Extraction yield increases with the increase of one of the parameters in a certain range while the other two conditions remain unchanged. However, there are also problems of increased cost, power consumption and unsafe factors. Generally, the extracting effect is rather good at 50 degrees Celsius and 20MPa for 5h. Supercritical carbon dioxide extraction used to extract soybean phospholipids has significance for the industrial application and is an applicative technology with wide prospect as it has the advantages of simple, non solvent residue, safe and reliable and high purity product and it consists with the trend of current green chemical technology (Shi,

The crude phospholipids are subjected to derosination and dissolved in solvents and then passed through ultrafiltration film with certain pore size. Components of suitable sizes pass

Ultrafiltration lecithin introduced by ADM(Archer Daniels Midland Co.) which has the property of dry, easy to be mixed with other materials, high quality and high purity is produced by removing the glycerides in phospholipids by ultrafiltration. Ultrafiltration

phospholipids without acetone residue are obtained and weighed for packaging.

phospholipids (Liu & Yang et al., 2006; Liu & Feng et al., 2006; Liu, 2007).

time needed to settle the whole phospholipids in acetone solution.

phospholipids product with purity of over 95%.

**3.2.2 Supercritical carbon dioxide extraction**

with purity of about 90% (Liu & Ma, 2011).

the best use of the materials.

**3.2.3 Ultrafiltration purification**

through the membrane and are isolated.

2005).

Mixed fatty acids and mixed fatty acid ethyl ester are added as fluidity agents during the vacuum concentrating procedure to improve the fluidic property of concentrated phospholipids and prevent phospholipids separating with the oil and guarantee the stability of phospholipids products.

The products obtained can flow freely at room temperature. If mixed fatty acids added is inadequate, it will not act as fluidity agent. On the contrary, excess addition of mixed fatty acids may raise the AV of phospholipids and get them rancid. The amount of mixed fatty acids added is usually 2.5%-3% (w/w) of the concentrated phospholipids. The addition of mixed fatty acid ethyl ester does not affect the AV and flavor of phospholipids and can gain high quality products but the cost is high. The amount is 3%-5% (w/w) of the concentrated phospholipids (Ji & Li, 2005).


Visual Cortex Current Status and Perspectives

*Edited by Stéphane Molotchnikoff and Jean Rouat*

**VISUAL CORTEX –**

**CURRENT STATUS**

and **Jean Rouat**

**AND PERSPECTIVES**

Edited by **Stéphane Molotchnikoff**


**1. Introduction**

Biodiesel is renewable, biodegradable and non-toxic transport fuel composed of fatty acid methyl esters. It is produced by transesterification of triacylglycerols with alcohol (mostly methanol and ethanol) in the presence of alkaline catalyst (e.g. sodium hydroxide). Common feedstocks for the production of biodiesel are different vegetable oils including rapeseed oil, palm oil, cooking oil, soybean oil and sunflower oil [1]. Production of biodiesel increased steadily in the last few years. In year 2016, the United States and Brazil were the world's biggest biodiesel producers with a production volume of around 5.5 and 3 billion liters, respectively [2]. All existing diesel engines, vehicles and infrastructure can run on pure biodiesel (B100) or blends with petroleum diesel fuel without any change [1]. Use of biodiesel has positive environmental impact, improves energy supply security, stimulates economic development and generates employment especially in the rural areas [3]. It reduces harmful emission characteristic for diesel exhaust such as particulate matter, carbon monoxide and total unburned hydrocarbons. Additionally, emission of toxic compounds including vapor-phase hydrocarbons from C1 to C12, aldehydes and ketones up to C8 and polyaromatic hydrocarbons and nitrated polyaromatic hydrocarbons are also decreased [4].

microorganism since they grow much faster than the microalgae. Unlike microalgae, they can use more diverse sugars and other carbon sources for their growth and lipid synthesis monosaccharides (glucose and xylose), amino sugars (N-acetylglucosamine), disaccharides (lactose, galactose, mannose, cellobiose and sucrose), alcohols (glycerol, ethanol and methanol), polysaccharides (starch and pectin) and organic acids (volatile fatty acids and acetic

In order to increase economic feasibility of the process production, different low-cost substrates have been used for the production such as crude glycerol, lignocellulose biomass (hydrolysate sweet sorghum bagasse, rice straw hydrolysate and corn stover hydrolysate), molasses waste, waste streams from food industry (whey permeate, olive pomace oil and olive oil mill wastewaters) and waste spent yeast from the brewing industry [5, 25–31].

The fatty acid biosynthetic pathway in most of oleaginous microorganisms is similar to nonoleaginous microorganisms. Two features of oleaginous microorganisms make them an efficient producer of lipid such as ability to efficiently produce precursor acetyl-CoA and cofactor NADPH needed for fatty acid synthesis. Process of lipogenesis could be divided in two steps, synthesis of precursor acetyl-CoA followed by biosynthesis of triacylglycerols (**Figure 1**). Nitrogen starvation in the presence of excess of carbon sources triggers *de nov*o synthesis of lipids in oleaginous microorganisms. Depletion of other media component like phosphorus or sulfur can efficiently induce lipogenesis [16]. Exhaustion of the nitrogen source induces a series of the consecutive biochemical reaction in the cell. The activity of AMP deaminase (AMPD) is upregulated. It cleaves the AMP to inosine monophosphate (IMP) and ammonia ions that cell can use as a nitrogen source. Consequently, concentration of AMP is reduced

regulated. This enzyme in oleaginous microorganisms is allosterically regulated by its activator AMP. Isocitrate accumulates in mitochondria and isomerized to citrate by aconitase (A). Accumulated citrate is transported into the cytoplasm in exchange for malate (citrate/malate translocase, CMT). In the cytoplasm, ATP citrate lyase (found only in oleaginous microorganisms) converts citrate to acetyl-CoA and oxaloacetate [32]. The acetyl-CoA is used for fatty acid synthesis while oxaloacetate is converted to malate by malate dehydrogenase (MDH)

The synthesis of lipids depends on efficient supply of NADPH, which is used for acetyl group reduction in the growing acyl chain. For the synthesis of 1 mol of a C18 fatty acid, 16 mol of NADPH is required. There is no unique metabolic route for generating NADPH in the oleaginous microorganism. Ratledge [33] described several routes for the synthesis of NADPH in the cytosol during lipogenesis. Transhydrogenase cycle which includes three enzymes pyruvate carboxylase (PC), MDH and malic enzyme (ME) has been proposed as a major route for the NADH production in the oleaginous microorganism. However, presence of ME in cytosol was not confirmed in some yeast species [16, 33]. In yeast *Y. lipolityca,* ME is located in the mitochondria and therefore cannot participate in the lipid synthesis [34].

)-depended isocitrate dehydrogenase (ICDH) is down-

Production of Microbial Lipids from Lignocellulosic Biomass

http://dx.doi.org/10.5772/intechopen.74013

139

acid) [5, 8, 20–24].

and the activity of NAD+

**2. Biochemistry of lipid accumulation**

(NADP+

and exported to mitochondria *via* CMT [16, 32].

Microbial lipids are viewed as an alternative feedstock for the biodiesel production because fatty acid compositions of accumulated lipids are similar to vegetable oils currently used as feedstock for the production of first generation biodiesel. Microbial lipids are also known as single cell oils (SCO), and are produced by heterogeneous group of oleaginous microorganisms that include less than hundred species of different microbial species including yeasts, fungi, bacteria and algae [3, 4]. Oleaginous microorganisms have the ability to accumulate significant amounts of intracellular lipids (more than 20% of their dry weight), mostly triacylglycerols, under certain cultivation conditions. Yeast strains such as *Lipomyces starkeyi, Rhodosporidium toruloides, Rhodotorula graminis, Rhodotorula glacialis* and *Trichosporon oleaginosus* can accumulate intracellular lipids from 50 to 80% (w/w) under certain cultivated conditions [5–8]. The fatty acid composition of lipids depends on the microbial strain and the cultivation conditions used. The most common fatty acids are palmitic (C16:0), stearic (C18:0), oleic (C18:1) and linoleic (C18:2) acids [9]. Microbial lipids of some oleaginous microorganisms are valuable source of polyunsaturated fatty acids that are used as additive for different food products and in nutriceuticals. Following omega-3 and omega-6 fatty acids are commercially produced using different wild-type and genetically modified oleaginous microorganisms such as γ-linolenic acid (GLA, C18:3, n−6) by *Mucor circinelloides*; dihomo-gamma-linoleic (DGLA) (20:3, n−6) by *Mortierella alpina* 1S-4; eicosapentaenoic acid *(*EPA) (20:5, n−3) by *Mortierella alpina* ST1358 and *Yarrowia lipolytica*; docosahexaenoic acid (DHA, 22:6, n−3) by *Crypthecodinium cohnii*, *Schizochytrium* and *Ulkenia* sp. and arachidonic acid (ARA, 20:4, n−6) by *Mortierella alpina* [10–17]. Microbial lipids from oleaginous yeast strains can be used as substitute for cocoa-butter and shea butter [18]. In comparison to vegetable oils, biodiesel production from microbial lipids have a number of advantages such as heterotrophic oleaginous microorganism grow much faster than the terrestrial crops; no need for arable land for cultivation; growth as well as cultivation does not depend on whether conditions and elimination of conflict between food and food supply chain [19]. Yeasts and fungi are favored oleaginous microorganism since they grow much faster than the microalgae. Unlike microalgae, they can use more diverse sugars and other carbon sources for their growth and lipid synthesis monosaccharides (glucose and xylose), amino sugars (N-acetylglucosamine), disaccharides (lactose, galactose, mannose, cellobiose and sucrose), alcohols (glycerol, ethanol and methanol), polysaccharides (starch and pectin) and organic acids (volatile fatty acids and acetic acid) [5, 8, 20–24].

In order to increase economic feasibility of the process production, different low-cost substrates have been used for the production such as crude glycerol, lignocellulose biomass (hydrolysate sweet sorghum bagasse, rice straw hydrolysate and corn stover hydrolysate), molasses waste, waste streams from food industry (whey permeate, olive pomace oil and olive oil mill wastewaters) and waste spent yeast from the brewing industry [5, 25–31].


**Slobodan P. Simonovic**

Department of Civil and Environmental Engineering, University of Western Ontario, London, ON N6A 3K7, Canada<EMAIL_ADDRESS>
Received: 20 September 2019; Accepted: 25 September 2019; Published: 27 September 2019

This book came as a support of the UNESCO International Hydrological Programme (IHP) activities on the International Shared Aquifer Resources Management (ISARM) project launched in the year 2000, with the goal of developing wise practices and guidance tools for the shared management of groundwater resources and to contribute to the multifaceted efforts required for global water cooperation. A key result of the project was the publication by UNESCO of the first world map of 592 transboundary aquifers. In spite of progress made, UNESCO's work required continuation. The book written by Jack Ganoulis and Jean Fried, with both authors being long-term collaborators of the UNESCO IHP, comes as a significant contribution to the transboundary groundwater education and training, offering a bridge between theory and practice.

The objectives set by the authors for the book included addressing the very important question of "water security" in an era of "major systemic risk" for humanity. The book starts with an old task of balancing water availability and water demand through "hydro-governance", defined as "an interactive process for managing at different levels and with different actors all kinds of water, including not only natural water resources but also new-water from wastewater after recycling and seawater desalination" (page vii). The book is correctly focusing on "transboundary hydro-governance" of water transboundary catchments that cover almost half of the world's land surface, including about 60% of global river flow and an area inhabited by 40% of the world's population. The main innovation introduced in the book is "integrated transboundary hydro-governance", which includes both surface water and groundwater. The book provides methodologies, practical tools, and examples of "effective transboundary hydro-governance".

The book is divided into three parts and eight chapters, starting with the main characteristics of transboundary waters (Part I), followed by the definition of transboundary hydro-governance (Part II), and ending with the discussion of transboundary hydro-governance in practice (Part III).

The first part offers (in three chapters) basic information on transboundary waters, water security, and topics of water conflicts and cooperation. The justification for employing a shared approach is clearly presented, and useful recommendations are provided for collaborative scientific approaches. Water security is also analyzed in this part, starting from the water variability in space and time. The ending chapter of Part I suggests the recognition of water districts at risk of conflict and the assessment and classification of different causes and various types of transboundary conflicts. The ending section focuses on the investigation of conflict prevention strategies and the ways to reverse potential conflicts into cooperation.

The second part of the book (in three chapters) provides clear distinctions between management, policy, and governance and analyzes some basic tools used to implement these concepts. This part is a

bit more technical. The main innovation presented in this section of the book is a multi-disciplinary integrated approach and its adoption to international settings. Discussion extends to an examination of what international joint institutions can be established and instruments that are needed to implement transboundary hydro-governance in practice. Tools and instruments (economic, shared, legal, and diplomatic) are reviewed in the last chapter of the Part II.

The third part of the book (in two chapters) is devoted to examples of hydro-governance in practice. Organizations and examples are studied from all around the world. From Danube to Mekong, and from Rhine to Senegal, the authors systematically analyze the pros and cons of approaches that lead to "good" transboundary hydro-governance. The analytical model of good hydro-governance can be derived from the experience presented in the book. However, in agreement with the authors and based on my own personal experience, there is still a long way to go.

This book is timely and pragmatically addresses worldwide problems, like climate change and significant floods and droughts, especially in parts of the world where every drop of water counts. The book can be used by experts and practitioners. Examples from the book could serve university courses on water resources management. The book has the potential to be a guiding manual and a reference tool.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The author declares no conflict of interest.

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