Journal of Medical Microbiology and Diagnosis-Biotechnology Open Access Journals

Mini Review

Oil Bodies as Novel Astaxanthin Carrier and Potential Aquafeed Ingredient for Aquaculture

Francisca Acevedo^1,2, Mauricio Quiroz³, Robert Simpfendörfer⁴, Mónica Rubilar^1,5 and Gonzalo Gajardo³

¹ Scientific and Technological Bioresource Nucleus, BIOREN, Universidad de La Frontera, Chile

² Department of Basic Sciences, Faculty of Medicine, Universidad de La Frontera, Chile

³ Laboratorio de Genética, Acuicultura & Biodiversidad, Universidad de Los Lagos, Chile

⁴Laboratorio de Bioquímica de Organismos Acuáticos, Universidad de Los Lagos, Chile

⁵Department of Chemical Engineering, Faculty of Engineering and Sciences, Universidad de La Frontera, Chile

Submission: February 21, 2018; Published: June 07, 2018

*Corresponding author: Acevedo F, Universidad de La Frontera, Casilla 54-D, Temuco, Chile.

Gajardo G, Laboratorio de Genética, Acuicultura & Biodiversidad, Universidad de Los Lagos, Av. Fuchslocher 1305, Osorno, Chile

How to cite this article: Francisca A, Mauricio Q, Robert S, Mónica R, Gonzalo G. Oil Bodies as Novel Astaxanthin Carrier and Potential Aquafeed Ingredient for Aquaculture. Adv Biotech & Micro. 2018; 9(4): 555768. DOI: 10.19080/AIBM.2018.09.555768

Abstract

This study presents preliminary evaluations for the potential use of Oil Bodies (OBs), spherical intracellular organelles that store lipids in the rapeseed Brassica napus, as a dietary supplement in aquaculture feeds. Objectives are straightforward: to evaluate OBs capacity to microencapsulate the highly valued but liable lipid-soluble pigment astaxanthin, used as feed supplement for appropriate coloration and to promote the health of aquatic animals. Likewise, to test the ability of Artemia, the biological model for micronutrient encapsulation, to safely uptake OBs and maintain astaxanthin stability over time in different Artemia bioencapsulation trials. Finally, to evaluate OBs qualitative lipid contribution. Oil bodies were extracted from rapeseeds with over 99% astaxanthin encapsulation efficiency. Astaxanthin was already visible one hourafter being encapsulated by Artemia, peaked at 24 hours, and remained non-metabolized into canthaxanthin for 48 hours. Significant differences were observed between the control group encapsulating astaxanthin-free OBs, and experimental groups, i.e., OBs enriched with four astaxanthin concentrations. Basal lipid content of Artemia increased qualitatively over time. It is concluded that OBs offer promise as a renewable and cost-effective agricultural carrier of astaxanthin and as alipid sourceto be potentially transferred to fish larvae via the live diet Artemia.

Graphical Abstract

A. Oil bodies (OBs) extracted from rapeseeds B. Astaxanthin microencapsulated in OBs (AST-M)

B. Qualitative assessment of lipid content in the gut of Artemia nauplii (Oil Red O stain), the biological model of encapsulation and live diet to transfer nutraceutics to fish larvae. Left, control. Right Oil Bodies ingested.

C. Bioencapsulation trials to track OBs ingestion over time, lipid contribution in Artemia nauplii and astaxanthin stability (not shown).

Keywords: Oil bodies; Rapeseed; Astaxanthin; Microencapsulation; Artemia

Abbrevations: OBs: Oil Bodies; AST: Astaxanthin; AST-M: Astaxanthin Microencapsulated within Oil Bodies

Introduction

Feedstuffs are finite and expensive ingredients which limits aquaculture diversification and expansion worldwide [1]. Fish oil represents a highly demanded but limited source of essential fatty acids used for dietary supplements in larval rearing (larviculture) of aquaculture species in captivity. Likewise, astaxanthin (AST) and other similar carotenoid pigments are valued but scarce feed supplements which are required for appropriate coloration and as a potent antioxidant to improve health status, since aquatic species exhibit poor ability to synthesize astaxanthin de novo. In the marine food chain, astaxanthin is primarily biosynthesized in microalgae and transferred to zooplankton and fish grazing on microalgae [2]. During marine plankton larvi culture, the natural diet of marine fish larvae is conveniently replaced by the microcrustacean Artemia whose nauplii (free swimming stages) are the live diet of choice during the critical period of metamorphosis, when marine fish larvae need to overcome radical molecular, physiological and morphological changes [3,4]. As a non-natural diet, Artemia often does not meet the nutritional requirements of fish larvae, and so a variety of boosters, or Artemia enrichment products are traded in the aquaculture market [5].

These products are bio-encapsulated in Artemia nauplii thanks to their ability to filter particles of less than 30μm. Lipid emulsions often produce such particle sizes when mixed with water, and these oil droplets are rich inessential fatty acids (EFA) such as eicosapentaenoic acid (EPA, 20:5 n-3), docosa hexaenoic acid (DHA, 22:6 n-3) and arachidonic acid (ARA, 20:4n-6), or a combination of these [3,6-8], depending on the booster used and bioencapsulation conditions [9]. Alternative micro diets like liposomes containing pigments, vitamins and free amino acids are also traded in the aquaculture market [10]. However, they are costly and short-lived as lipids oxidize easily. Additionally, the use of these boosters is unsustainable because they depend on fish oil which is increasingly scarce. Therefore, alternative plant oil sources have gained interest as they are globally available, inexpensive and can at least partially satisfy requirements offish larva [11,12].

Plant oils are generally rich in n-6 and n-9 types, mainly linoleic acid (18:2n-6) and oleic acid (18:1n-9), with moderate to low levels of n-3 such as α-linolenic acid (18:3n-3) (except linseed oil) [13]. With some limitations, rapeseed oils potentially emerge as suitable microdiets, or as potentially safe carriers of liable dietary supplements [14]. In oleaginous plants like rape seed (Brassica napus), oils are generally stored in spherical intracellular organelles referred to as oil bodies (OBs). They consist of an oil core with a matrix of triacylglycerol (TAG) bound by a phospholipid monolayer embedded with proteins known as oleosins [15]. Moreover, tocopherol and polyphenols are reported to be intrinsically bound to OB structures in Brassica napus seeds [16]. Therefore, OBs may be considered a natural protection system against fatty acid oxidation because they can safely store lipids in seeds, in the form of TAGs, for long periods. Due to their spherical shape and small size (0.5 to 2.0μm in diameter), OBs are considered the natural equivalent of liposomes [17], and may be useful for the development of safe and efficient carriers to deliver bioactive molecules for food and/or pharmaceutical purposes.

The aims of this study were to evaluate the ability of OBs to microencapsulate and safely carry the highly valued but liable lipid-soluble pigment astaxanthin, used as feed supplement for appropriate coloration and to promote health inaquatic animals. Likewise, to test the ability of Artemia, the biological model for micronutrient encapsulation, to safely uptake OBs and maintain astaxanthin stability over time in different bioencapsulation trials. Finally, to evaluate OBs qualitative lipid contribution.

Materials and Methods

Preparation of astaxanthin microencapsulated within oil bodies

Isolation of OBs from Brassica napus seeds: Mature Brassica napus seeds obtained from local producers (Gorbea, La Araucanía Region, Chile) were dried at 30 °C for 24h. Isolation was done following methods of Chen et al. [18] with slight modifications. Briefly, 50g of seeds were ground in a blender (Moulinex Super 96 Junior “S”); the sample was then mixed with 250mL of buffer solution (3mM concentration of MgCl2 and 100mM Tris-HCl, pH 8.5) and homogenized (200rpm, 3h, 4 °C, orbital shaker). The homogenate was filtered through three layers of Miracloth and the remaining filter cake was pressed. The collected slurry was then centrifuged (10,000×g, 30min, 4 °C) twice. The upper layer (“cream layer”) was isolated using a chilled metal spatula. The resulting slurry was filtered and centrifuged with what was collected after the first extraction step at 10,000×g for 30min at 4 °C. The cream layer containing crude oil bodies was collected and re-suspended (10% w/w in buffer solution 10mM Tris-HCl pH 8.5 containing sodium azide 0.02M) and stored at 4 °C until further use. Sodium azide was added to prevent microbial spoilage of the OB suspension.

Astaxanthin microencapsulation within oil bodies

Different astaxanthin solutions dissolved in dimethylsulfoxide were prepared (250, 500 and 1000ppm) according to Acevedo et al. [19]. An aliquot (10mL) of each solution was mixed with 10mL of OB suspension, and stirred at 200rpm for 5.3h. The orange cream layer containing AST microencapsulated or loaded within OBs (AST-M) was collected and washed twice with buffer solution (10mMTris-HCl pH 8.6). After microencapsulation, samples were centrifuged (6,793×g, 15min) for phase separation as described by Acevedo et al. [19]; the orange cream layer (AST-M) was recovered using a chilled metal spatulaand stored at 4 °C until further use [20-23].

OBs bioencapsulation by Artemia nauplii

Artemia cysts from Great Salt Lake (GSL, USA) were decapsulated and hatched according to the standard protocol by Sorgeloos et al. [24]. nauplii were collected and distributed (~180,000 nauplii) in five 250mL conical tanks with 250mL of artificial seawater (35ppt) at 25 °C. Each tank was illuminated and subject to high aeration in order to homogenize nauplii distribution in the water column. Experimental groups (three tanks) consisted of Artemia nauplii fed OBs loaded with different astaxanthin concentrations (AST-M: 250, 500 or 1000ppm). For this, aliquots (1.8mL) from a 10% w/w OBs suspension were added in a single dose at the start of the bioencapsulation experiment. Control groups (2 tanks) corresponded to starved nauplii and a group fed free AST-M. Independent samples (1.0mL each) were taken at 1, 6, 12, 24, 48 and 72 hours for analysis: two for fluorescence microscopic analysis (fixed in 25% v/v acetic acid), and two for qualitative lipid staining (fixed in 4% v/v formalin) by means of the Oil Red Olipid staining technique. Fixed samples were stored at 4 °C for further analysis.

**Quantification of astaxanthin concentration in Artemia nauplii by HPLC**

Five groups of Artemia nauplii with three replicates each were independently compared: i) control (starved nauplii); ii) nauplii fed OB suspension (10% w/w); iii) nauplii enriched with AST-M: 250; 500 and 1000ppm. A single OB dose (1.5mL in 250mL artificial seawater) was provided for enrichment and temporal monitoring was done in two groups, one tested at 1, 6 and 12h, and the other at 24 and 48h. nauplii mortality was visually checked to avoid manipulating the enrichment process, and to keep a reasonable number of nauplii for analysis. Mortality in the experimental groups was insignificant compared to the starved control, in which dead nauplii sank to the bottom of the tank. nauplii samples (20mg wet weight) from each group were taken for HPLC analysis at 0,1,6,12,24 and 48 hours. Time zero corresponded to nauplii sampled at the beginning of the enrichment experiments (24h post-hatch). The collected nauplii were thoroughly washed with distilled water in a mesh net to remove salt and extra-corporal OBs, then weighed and stored at -20 °C until HPLC analysis. nauplii biomass was completely homogenized in 350μL of acetone using a glass homogenizer. Samples were centrifuged for 1min at 13,000rpm, and 20μL of each organic extract was immediately analyzed by HPLC-DAD. The RP-column was from Kromasil (C-8, 50 x 4.6mM, 2.5μm particle size). The high-performance liquid chromatograph used was from Jasco (Japan), with a quaternary pump (PU-2089 Plus model) and a diode array detector (MD-2015 Plus model). Chrompass software (Jasco) was used for the chromatogram analysis. Solvents used to develop the gradient were (A) Milli-Q water and (B) methanol. Solvent gradient in volumetric ratios for solvents A and B was as follows: 0-5.6min, 12A/ 88B; 5.6-7.6min, 0A/100B; 7.6-7.8min, 0A/100B; 7.8-9.5min, 12A/88B. Astaxanthin and canthaxanthin carotenoids were detected at 478nm absorption, and quantified by comparing the peak areas with the respective calibration curves (external standards, both from Sigma Chem. Co.). No internal standard was used because the extraction procedure was quantitative, reproducible, and required only one extraction step. Retention times for AST and canthaxanthin were 3.13min and 5.65min, respectively. Flow rate was set to 0.7mL/min.

Qualitative lipid assessment

Triglyceride-specific Oil Red O histological staining [20] was used to visualize lipid presence in the gut of Artemia nauplii. Formalin-fixed samples were rinsed with 60% v/v isopropanol and centrifuged at 12,000 rpm for 2min. Samples were stained with freshly prepared Oil Red O (0.3% v/v in isopropanol) working solution for 20min, and then rinsed with 60% v/v isopropanol and distilled water. After centrifugation at 12,000rpm for 2min, samples were stained with violet crystal (contrast), diluted twice and rinsed with 60% v/v isopropanol and distilled water. Finally, samples were suspended with 100μL distilled water. Image J 1.48v software was used to quantify the stained area of Artemia nauplii images (http://imagej.nih.gov/ij/docs/examples/stainedsections/ index.html). Average (±SD; 10 individual nauplii) ratio of the stained area vs. total area was obtained for each group. The median image was taken from each group (Image/Stacks/Images to stack) using “Z Project”.

Statistical significance of AST incorporated in Artemia nauplii was analyzed by a multivariate ANOVA using the “R” software, with Tukey’s post-test. Four diets (AST free-OBs and OBs enriched with different astaxanthin concentration, AST-M diets) and five enrichment times (1, 6, 12, 24 and 48h) were compared. Lipid accumulation-related differences were analyzed using oneway ANOVA. Differences between means were compared using Duncan’s test. A significance level of 95% (p<0.05) was used throughout.

Result and Discussion

Oil bodies were extracted from Brassica napus seeds with a yield of about 30% (50gr of rapeseed initially grinded). Figure 1 shows selected OBs loaded with astaxanthin (AST-M). The encapsulation efficiency of AST with OBs was over 99% as determined by the methodology of Acevedo et al. [19]. In a scenario of scarce basic aquafeed ingredients, the aquaculture industry has switched to plants as more sustainable alternative sources [1]. This study showed some practical advantages of OBs such as easy and cheap extraction, as well as being a renewable and available agricultural product [19]. Additionally, OBs were easily enriched with different astaxanthin concentrations. As hydrophobic molecule, astaxanthin exhibits high affinity for the triacylglycerol core of OBs which facilitates its microencapsulation by passive diffusion in a simple, fast (less than 6 hours) and efficient process, as previously shown (over 99% encapsulation efficiency) [19].

Figure 2A illustrates the results of Artemia bioencapsulation experiments in which OBs loaded with different astaxanthin concentrations (AST-M) were monitored over time in Artemianauplii. Astaxanthin was visible in all groups except control (starved nauplii), one hour after encapsulation experiments began, and peaked at 24hours in all experimental groups. Astaxanthin concentration was higher in Artemia nauplii taking up OBs with higher astaxanthin concentration. Statistically significant (p<0.05) differences were observed between control, bioencapsulating AST-free OBs, and the AST-M experimental groups (with 250, 500, 1000ppm) (Figure 2A). No statistically significant (p>0.05) differences were observed in the Artemiaendogenous content of canthaxanthin (Figure 2B) according to the diet provided, implying that astaxanthin was not metabolized into canthaxanthin under these study conditions. Optimal AST microencapsulation time was 24hours.

An initial concern was OB size (average diameter 0.5 to 2.0μm), which is below the range of 7-28μm filtered by Artemia nauplii [21], with an optimum around 16μm. This study made it clear that Artemia nauplii can ingest particles below the pre-established size limit, though bioencapsulation efficiency depends on several experimental conditions [9]. Astaxanthin stability was tested for at least 48hours in Artemia which confirmed the hypothesisthat OBs are safe astaxanthin carriers [19]. This is not normally the case with emulsions used to boost Artemia, which tend to be quickly digested, and explains why the bio encapsulation process is often relatively fast [22]. Thus, an extra benefit of bio encapsulating AST-M is that, unlike unprotected supplements, enriched nauplii can be kept stable for relatively longer periods without affecting biological properties of the ingested substance. By extending the pre-established window time for efficient Artemia bio encapsulation (around 24 hours), OBs provide a chance for nauplii to grow beyond the size normally reached at 24hours, and hence be available for larger fish larvae (greater mouth opening). Protection provided by OBs would explain why astaxanthin was not metabolized into can thaxanthin, the most abundant carotenoid in Artemia. Isomeric canthaxanthin forms (cis and trans) vary during the Artemia life cycle, depending on the availability of its metabolic precursors: echinenone and betacarotene [23].

Figure 3 shows Oil Red O staining for Artemia nauplii individuals after 1 to 72 hours post encapsulation for the following groups: 1) control group (starved nauplii) in which stain intensity decayed over time; 2) nauplii bioencapsulated with AST-free OBs, which exhibited higher intensity and 3) nauplii that bio encapsulated OBs loaded with different astaxanthin concentrations. As expected, there are no clear visual differences between groups 2 and 3 since the Oil Red O stain does not show differences in AST concentration. Image quantification shows the ratio of stained area vs total area (in percent) increased as bioencapsulation proceeded, from 6.0% at 24 hours in the control, 10.1% in the group fed OBs and 13.9 and 13.7% in those nauplii fed OBs with increasing astaxanthin concentration (500 and 1000ppm). This means that nauplii reached the maximum stain intensity at 24hours, which is a proxy of the basal lipid quantity and/or OBs concentration in the gut of Artemia nauplii.

Testing the safe OBs uptake by Artemia nauplii is a precondition to transfer AST to marine fish larvae, owing to the role this microcrustacean plays in the food chain of animals reared in captivity. However, Oil Bodies enriched with astaxanthin could be directly provided to fish larvae, or even to shellfish larvae that are also boosted with dietary supplements [24]. The advantage of OBs as a lipid source would be that they are already naturally emulsified after the extraction, with remarkable physical and chemical stability and potentially high nutritional value, depending on the species targeted [25].

Conclusion

A. OBs are cheap and renewable agriculture products that can potentially be used as microdiets to safely carry lipidsoluble pigments like AST, which is highly valued for aquatic species due to its nutritive and biological properties. In this study OBs were efficiently microencapsulated with different astaxanthin concentrations [26].

B. OBs were encapsulated by Artemia nauplii, the biological model for encapsulation that is additionally used as live diet in the food chain of marine fish reared in captivity. However, OBs could also be delivered directly to fish or shellfish, in the latter case as a complementary or partial replacement for lipid emulsions.

C. Artemia nauplii ingested and stored OBs loaded with AST for over 24 hours, the operational time defined for optimal Artemia enrichment, thus extending the enrichment time window to allow larger fish larvae to feed on Artemia. Astaxanthin remained non-metabolized in the gut of Artemia nauplii for longer times, likely protected inside OBs lipid core [27].

D. OBs supplemented the basal lipid composition of Artemia nauplii, and since they are rich in a variety of lipids, they appear to be a naturally good provider of such an essential diet ingredient.

Funding

Authors report no conflict of interest, and they are solely responsible for the content and writing of this article. Authors acknowledge Esteban Camaaño for technical support with the Red Oil staining. Funding from the National Commission for Scientific and Technological Development (CONICYT) through projects FONDECYT3120022 (to FA) and FONDEF D09E1256 (to GG) is gratefully acknowledged.