Despite tremendous advances in medicine around the world, infectious diseases caused by bacteria, fungi, and different types of protozoa continue to pose a major threat to public health. Their impact is particularly significant in developing countries due to the lack of access to medicines and the emergence and spread of antibiotic resistance, which has increased dramatically over the past decade (Kang et al. 2015). Several antimicrobial chemical agents have been synthesized for certain parasitic and bacterial diseases, however, they are still very limited in number and their use is restricted due to their toxicity, high cost, microbial resistance and low efficacy (Sanchez et al. 2010). According to the Eastern Mediterranean Regional office of the World Health Organization, cutaneous leishmaniasis (CL) caused by the protozoan
On the other hand, oxidative damage caused by reactive oxygen species (ROS) on lipids, proteins, and nucleic acids can trigger various chronic diseases, such as coronary heart disease, atherosclerosis, cancer, and aging (Ngo et al. 2011). Over the past decades, the food, cosmetics, and nutraceutical industries have been putting enormous efforts into finding natural alternatives to replace synthetic antioxidants, in particular BHT (butylated hydroxytoluene) and BHA (butylated hydroxyanisole), which several epidemiological studies have shown to be toxic in the medium and long term (Agyei et al. 2015). Such alarming situation has highlighted the urgency of finding new safe alternatives of natural origin, easily accessible, and capable of being produced in a sustainable manner.
In recent years, the use of photosynthetic microorganisms such as microalgae in life sciences has received increasing attention due to their diverse phytometabolic content with varied chemical structures and biological properties. The group of phycobiliproteins (phycocyanin, allophycocyanin and phycoerythrin) are among the most valuable accessory pigments that can be extracted from microalgae due to their extremely beneficial effects on health. They are widely recognized as powerful functional ingredients with very interesting applications in various fields such as agri-food, aquaculture, biotechnology, medicine, and pharmacy (Lamers et al. 2010; Jalal et al. 2013).
Currently, the growing demand for functional foods and nutraceuticals has sparked an increasing interest in the use of microalgae as natural sources of high value products, especially phycobiliproteins, which has automatically resulted in their surge in demand in the global market of human health, nutrition, and aquaculture (Jalal et al. 2013; Yang et al. 2013). However, their wide industrial application still requires studies to isolate and characterize more candidate strains from various terrestrial and marine environments in order to select those with interesting levels of accumulation of these valuable molecules.
Therefore, the present work aimed to study the production of phycobiliproteins and the accumulation potential of some local microalgal strains isolated in several Tunisian inland water bodies, as well as to determine their phytochemical composition and to evaluate antioxidant, antibacterial, and antileishmanial properties of their extracts.
The microalgae strains investigated in this work were collected in eight Tunisian inland water bodies (river, lagoon, dam reservoir, spring) located in the north, center, and south of the country (Table 1).
Microalgal strains isolated from Tunisian inland water bodies (river, lagoon, dam reservoir, spring) with location and culture mediums
Strain | Class/Order | Sampling site | Culture medium | |
---|---|---|---|---|
Duna-GV | Chlorophyceae/Chlamydomonadales | El Grine Sabkha |
CONWAY | |
Chroo-CH | Cyanobacteria / Chroococcales | Chanchou River |
BG11 | |
Arthro-KB | Cyanobacteria / Oscillatoriales | Korba lagoon |
BG11 | |
Osci-NB-01 | Nabhena reservoir |
BG11 | ||
Lepto-CH | Chanchou River |
BG11 | ||
Osci-BM-01 | Bir Mcherga reservoir |
BG11 | ||
Plank-SS-01 | Sidi Saad reservoir |
BG11 | ||
Spir-ML | Maltine River |
CONWAY | ||
Pseud-01 | Cyanobacteria / Nostocales | Siliana spring |
BG11 | |
Cyl-NB-05 | Nabhena reservoir |
BG11 |
S – salinity; PSU – practical salinity unit
All samples were taken to the laboratory where each strain was subjected to capillary isolation in liquid medium, using an inverted microscope (Leica microsystems, Wetzlar, Germany). A single individual was isolated (a single cell, colony, or filament) in 300 μl of the appropriate culture medium, and growth of each isolate was achieved by gradually increasing the culture volume from the initial isolation to the final batch. All strains were cultivated in 2 l laboratory flasks, using a monospecific batch culture system under sterile conditions. Growing was carried out at 25 ± 1°C temperature in a thermostatically controlled room, and using BG11 (Rippka et al. 1979) or CONWAY (Blancheton 1985) media, depending on the origin of each strain (Table 1). Culture media were sterilized by autoclaving at 120°C for 20 min before use. Growth was conducted in a 16:8 h light:dark regime under illumination of cool white fluorescent tubes with a light intensity of approximately 25 μmol m−2 s−1. Cells were harvested only in the exponential growth phase by centrifugation (3000 rpm; Thermo IEC CL31R Multispeed centrifuge; Massachusetts – USA), then lyophilized using a Christ Alpha 2-4 LD Plus freeze-dryer (Harz, Germany) after a cultivation period between 18 and 25 days. Samples were then stored at −20°C until analysis.
For screening of the investigated biological activity, the lyophilized biomass of each strain was ground using a mortar and pestle for a few minutes and extracted adding 1:10 w/v of solvent mixture (dichloromethane–methanol, 1:1, v/v) with the assistance of an ultrasonic bath (Branson 2800, Branson Ultrasonics Corporation, Danbury, USA) (110 w; 40 kHz) for 15 min at room temperature, and then filtered. The filtrate was subsequently collected and the remaining microalgal material was ground in a mortar, mixed with the same extraction buffer and filtered again until the completion of three successive extraction cycles for maximum extraction yield. The solvents were then evaporated using a Rotavapor (KNF, Freiburg, Germany) and the obtained extracts were stored at −80°C between each analysis.
Morphology and size of the microalgae strains were observed through a Leica DM LS2 microscope (Leica microsystems, Wetzlar, Germany) with magnification: 400x and 1000x, using a specialized digital camera (HDCE - 50B, Olympus Corporation, Tokyo, Japan) monitored by AxioVision software, version 4.8. Taxonomic identification was based on Geitler (1932), Bourrelly (1966) and Komárek and Anagnostidis (1999, 2005).
Total genomic DNA was extracted using a DNeasy® Plant Mini Kit (QIAGEN GmbH, Hilden, Germany) following the protocol provided by the manufacturer and stored frozen at −20°C until use. The DNA extracts were checked by electrophoresis in 1% agarose gel and then quantified with a Thermo Fisher spectrophotometer (Heaios Y, Thermo Scientific; Massachusetts, USA), followed by dilution to 200 ng/μl solutions. Polymerase chain reaction (PCR) amplifications were performed with a DNA Thermal Cycler, model 2720 (Applied Biosystems/California, USA), using 100 μl of the PCR reaction mixture. For prokaryotic strains, nuclear-encoded 16S rRNA gene segments were amplified with PCR primers 27 Fw and 1494 Rev under the conditions described in Fathalli et al. (2011). For
All PCR amplifications were performed in 20 μl aliquots, containing 10 pmol of each forward and reverse primers (Invitrogen, Carlsbad, California), 1 × Reaction Buffer (Invitrogen, Carlsbad, California), 250 μM of each deoxynucleoside triphosphate (OMEGA biotec TQAC136), 2.5 mM MgCl2 (Invitrogen, Carlsbad, California), 0.5 U of Taq DNA polymerase (Invitrogen, Carlsbad, California), and 5 to 10 ng of genomic DNA template. All PCR amplicons were analyzed by electrophoresis in 1% agarose gel (Invitrogen, Carlsbad, California), run in 1 × TBE buffer, stained with ethidium bromide, and photographed under UV trans-illumination.
A total of 100 μl of PCR of each amplified product was purified using a PureLink® PCR Purification Kit (Invitrogen, Carlsbad, California), following the protocol provided by the manufacturer, and subsequently sent for direct sequencing. Nucleotide sequences were obtained and submitted to the BLAST (Basic Local Alignment Search Tool) database (
All sequences were then submitted to the GenBank database (accession numbers are given in Table 2).
Molecular identification of microalgal strains isolated from different Tunisian inland water bodies with % of similarity according to the BLAST database and accession numbers
Strain | Taxon according to BLAST database | % of similarity | Accession numbers | |
---|---|---|---|---|
16S rRNA | rpoC1 | |||
Osci-NB-01 | 99 | MG762090 | - | |
Osci-BM-01 | 100 | MG762091 | - | |
Plank-SS-01 | 100 | MG762092 | - | |
Cyl-NB-05 | 96 | - | HQ389355 | |
Pseud-01 | 99 | MG098078 | - | |
Arthro-KB | 100 | MG098079 | - | |
Lepto-CH | 99 | MG098077 | - | |
Spir-ML | 99 | MG518486 | - |
The content of phycobiliproteins (phycocyanin – PC, allophycocyanin – APC, and phycoerythrin – PE) was determined during the exponential growth phase of cyanobacterial strains according to Bennett & Bogorod (1973). For each strain, 10 ml of fresh culture, taken in a culture concentration range of 100 μg of fresh weight per ml, was centrifuged at 3000 rpm for 5 min. The collected cell mass was then washed with buffer solution 1 M Tris HCl (pH 8.1), and one volume of cell mass was subsequently resuspended in five times the volume of the same buffer. In order to extract the pigments, it was necessary to perform cell wall splitting for cyanobacterial strains. For this, continuous freezing at −20°C, thawing at +4°C, and sonication (10 min with cycles of 30 s) were applied to all samples, which allowed destruction of cell walls. The cell fragments were then separated by centrifugation at 12 000 rpm for 10 min, and the supernatant was taken for spectrophotometric determination of phycobiliprotein content by measuring the absorbance at wavelengths of 620, 652 and 562 nm. Absorbance measurements were performed on a UV/Vis spectrophotometer (Heaios Y, Thermo Scientific; Massachusetts, USA), and the amount of PC, APC, and PE in a sample was calculated using the following formula (Bennett & Bogorad 1973; Horvath et al. 2013):
Each sample was analyzed in triplicate and the buffer was used as a blank.
The total content of polyphenols (TPP) of each extract was assessed according to a colorimetric method adapted from Dewanto et al. (2002) using the Folin–Ciocalteu reagent. A volume of 0.125 ml of ethanol extracts was added to 0.5 ml of distilled water and 0.125 ml of Folin–Ciocalteu reagent. After 6 min incubation in the dark, the mixture was added to 1.25 ml of 7% (w/v) sodium carbonate, and the solution was made up to 3 ml with distilled water. The mixture was then incubated for 90 min in the dark and then taken for absorbance measurement at 760 nm, using a UV/Vis spectrophotometer (Optizen 2120 UV plus, Mecasys, Korea). TPP was calculated from the linear equation of a standard curve prepared with gallic acid (concentration range of 50–200 μg ml−1). Results were expressed in mg of gallic acid equivalents per gram of dry weight (mg GAE/g DW).
Total flavonoid (TF) content was determined colorimetrically according to Heimler et al. (2006). Briefly, 0.075 ml of 7% NaNO2 (w/v) was mixed with 0.25 ml of each extract and 0.15 ml of 10% (w/v) AlCl3. After 5 min incubation at room temperature, 0.5 ml of (1 M) NaOH was added to the mixture and adjusted to 3 ml with distilled water. Absorbance was determined at 510 nm after 15 min incubation at room temperature. The TF content was estimated based on a standard calibration curve (concentration range of 100–750 μg ml−1). Results were expressed in mg catechin equivalent per gram of dry weight (mg CE/g DW).
Antioxidant activity of microalgal extracts was assessed using 1,1 Diphenyl 2-Picryl Hydrazyl (DPPH) assay for free radical scavenging activity according to Cheel et al. (2007). Samples of microalgae extracts were prepared in methanol at a final concentration ranging from 62.5 μg ml−1 to 2 mg ml−1. They were subsequently mixed with 2 ml of (60 μM) DPPH solution prepared in methanol, then incubated in the dark for 30 min at room temperature. Absorbance measurements were taken against a blank at 517 nm using a UV/Vis spectrophotometer and butylated hydroxytoluene (BHT) as a standard. Free radical scavenging activity was determined according to the following equation:
IC50 was expressed as the concentration of extract inhibiting 50% of the DPPH free radical scavenging activity.
Antimicrobial activity of microalgal extracts was tested against reference microorganisms, including Gram-positive bacteria (
Antibacterial activity tests were carried out using the agar disc diffusion method according to Celiktas et al. (2007). Bacterial suspensions (108 cells ml−1) were inoculated evenly in Mueller-Hinton (MH) agar plates using a cotton swab. Whatman filter paper discs (diameter = 5 mm, porosity = 6 μm) were impregnated with double serial dilutions of the extract, ranging from 31.25 μg ml−1 to 1 mg ml−1, then allowed to dry under a flow hood for 3 h before application to the agar surface. Plates were incubated at 37°C for 24 h, and the size of inhibitory zones (including the diameter of the disc) was measured. Clear zones of inhibition around the discs indicate antibacterial activity. The minimum inhibitory concentration (MIC) was determined for extracts with a zone diameter over 10 mm. The MIC was determined on 96-well plates and defined as the lowest concentration of the extract that inhibits the growth of microorganisms. The tetracycline (30 μg/disk) was used as a reference antibiotic.
Antileishmanial activity was evaluated in 96-well plates on promastigotes of
All solvents and reagents used for the extraction procedures, antioxidant activity, and antileishmanial assay were purchased from Sigma-Aldrich (GmbH, Steinheim, Germany). Amphotericin B was purchased from Sigma (98% purity, Sigma–Aldrich, USA).
Assays were conducted in triplicate and data were expressed as the mean ± standard error of three independent measurements. To assess the significance of linear correlations between the antioxidant activity and total phenolic and flavonoid amounts in each microalgal extract, Pearson’s correlation analysis was carried out using r-Project software. Differences were considered significant at
Quantitative evaluation of PC, APC, and PE content was carried out on the cyanobacterial strains studied as shown in Fig. 1.
Phycobiliprotein concentration of cyanobacterial strains isolated from different aquatic habitats expressed in μg per ml of fresh culture. Bars indicate the standard deviation of three replicates (PC = Phycocyanin; APC = Allophycocyanin; PE = Phycoerythrin)
Nostocales cyanobacterium Cyl-NB-05 contained the maximum total phycobiliprotein content (3150 μg ml−1 of fresh culture), while the minimal amount was found in Chroo-CH and Pseud-01 (44 μg ml−1 each), within the same culture concentration range (~100 μg ml−1). For the top phycobiliprotein producer strain (
In terms of APC production, the maximum yield was found in Osci-NB-01 (362 μg ml−1), which was also the only isolate where the amount of APC largely exceeded its PC level (twice as much). It was closely followed by another Oscillatoriales cyanobacterium, Plank-SS-01 (326 μg ml−1).
In all the strains studied, PE was present in lower amounts compared to the two other constituents, with the exception of Chroo-CH and Pseud-01, where it was the major phycobiliprotein produced.
The amounts of total polyphenols and flavonoids (TPP and TF, respectively) contained in the crude extracts of the investigated isolates are shown in Table 3. TPP levels varied between undetectable (
Phenolics, flavonoids and antioxidant activity of microalgal extracts
Strain | Code | TPP (mg EqGA/g DW) | TF (mg EqC/g DW) | Antioxidant activity against DPPH (IC50, μg ml−1) |
---|---|---|---|---|
Chroo-CH | 8.90 ± 0.25 | 67.94 ± 1.36 | 212.15 ± 2.07 | |
Osci-NB-01 | 1.81 ± 0.39 | 58.46 ± 0.73 | 263.91 ± 14.93 | |
Arthro-KB | 2.85 ± 0.06 | 74.09 ± 5.28 | 953.94 ± 28.14 | |
Lepto-CH | 7.75 ± 0.77 | 68.17 ± 3.96 | 782.87 ± 4.2 | |
Plank-SS-01 | 4.86 ± 0.3 | 38.32 ± 5.37 | NA | |
Pseud-01 | 5.63 ± 0.15 | 38.05 ± 4.43 | 912.21 ± 2.21 | |
Cyl-NB-05 | ND | 1.48 ± 0.03 | NA | |
Osci-BM-01 | 1.77 ± 0.13 | 36.31 ± 3.12 | NA | |
Spir-ML | ND | 5.03 ± 1 | NA | |
Duna-GV | 3.07 ± 0.32 | 37.94 ± 3.11 | 681 ± 20,07 | |
BHT | - | - | - | 17.34 ± 0.23 |
TPP – total polyphenols; TF – total flavonoids; DW – dry weight; GA – gallic acid ; C – catechin ; BHT – butylated hydroxytoluene ; NA – not active at maximum concentration; ND – not detectable. Results represent means of three replicates with standard deviation (Means ± SD, n = 3)
The antioxidant activity of microalgal extracts against DPPH free radicals are shown in Table 3. The higher the DPPH radical scavenging activity of a compound, the lower its IC50 value (inversely proportional), since the IC50 represents the amount of antioxidants needed to reduce the concentration of a free radical by 50%.
The lowest IC50 values were found in
The antibacterial activity of microalgae extracts was studied against human pathogenic bacteria (Table 4). Most strains of microalgae showed no detectable antibacterial activity against all species of bacteria tested, except for
Antibacterial activity of microalgal extracts
Strain | Inhibition zone diameter (mm) [MIC (μg ml−1)] | |||||
---|---|---|---|---|---|---|
MRSA | ||||||
Chroo-CH | NA | NA | NA | NA | NA | NA |
Osci-NB-01 | NA | NA | NA | NA | NA | NA |
Arthro-KB | NA | NA | NA | NA | NA | NA |
Lepto-CH | NA | NA | NA | NA | NA | NA |
Plank-SS-01 | NA | NA | NA | NA | NA | NA |
Pseud-01 | NA | NA | NA | NA | NA | NA |
Cyl-NB-05 | NA | NA | NA | NA | NA | NA |
Osci-BM-01 | NA | NA | NA | NA | NA | NA |
Spir-ML | NA | NA | NA | NA | NA | NA |
Duna-GV | NA | NA | NA | 16 ± 0.1 [375] | NA | NA |
NA – not active at maximum concentration
The antileishmanial capacity of microalgae extracts was tested against the promastigotes
According to the results obtained, the
Antileishmanial activity of microalgal extracts
Strain | IC50 ± SD (μg ml−1) | |
---|---|---|
Chroo-CH | 713.4 ± 0.24 | 1000 ± 1.96 |
Osci-NB-01 | 448.7 ± 0.24 | 728.5 ± 1.21 |
Arthro-KB | > 1000 | > 1000 |
Lepto-CH | 212.7 ± 1.46 | 401.9 ± 1.22 |
Plank-SS-01 | 1000 ± 1.98 | > 1000 |
Pseud-01 | 1000 ± 2.38 | > 1000 |
Cyl-NB-05 | 246.4 ± 1.66 | 473.8 ± 0.53 |
Osci-BM-01 | 188.7 ± 1.35 | 377.5 ± 0.09 |
Spir-ML | 1000 ± 2.44 | > 1000 |
Duna-GV | 151.2 ± 0.37 | 284.87 ± 1.46 |
Amphotericin B | 0.64 ± 0.24 | 0.97 ± 0.08 |
IC50 = Inhibitory Concentration 50 (μg ml−1); Amphotericin B = positive control.
SD – standard deviation; each value was represented as mean ± SD (n = 3)
To the best of our knowledge, this work is the first to highlight the capacity of the cyanobacterium
In addition to the previously described beneficial effects, some epidemiological studies have demonstrated the efficacy of PC as an inhibitor of cancer cell proliferation
Moreover, the neuroprotective role of PC against global cerebral ischemia/reperfusion damage in gerbils has been reported (Penton-Rol et al. 2011). PC was found to exhibit a potent protective effect against neuronal cell death in the hippocampus, and significantly improve locomotor behavior and survival in gerbils after just one week of reperfusion. In another study, Rimbau et al. (2001) found strong evidence for a remarkable protective role of PC against cell death caused by 24 h withdrawal of potassium and serum (K/S) in rat granular cerebellar cell cultures. These findings highlight some promising perspectives on the use of our
When comparing our results with previous studies, it can be observed that, in general, the levels of antioxidant activity against DPPH radicals obtained in the present work remain relatively less pronounced. IC50 values ranging from 67.5 to 119.6 μg ml−1 and from 30.72 to 102.47 μg ml−1 were reported by Ijaz & Hasnain (2016) and Babic et al. (2016), respectively, for the antioxidant activity of crud extracts of several cyanobacterial strains against DPPH radicals. Similarly, Safafar et al. (2015) recorded 10-fold higher inhibitory activity for
The particularly interesting antioxidant activity recorded in
This finding appears to be consistent with previous studies, e.g. Blagojevic et al. (2018), where an extremely low IC50 value was reported for BHT (about 10 μg ml−1), reflecting much stronger DPPH scavenging efficiency compared to 10 microalgal isolates essayed in their study.
Nevertheless, it should be noted that such extraordinary antioxidant efficiency of BHT among other synthetic commercial antioxidants, especially butylated hydroxyanisole (BHA), tertbutyl hydroquinone (TBHQ), or propyl gallate (PG), is not without a significant cost. In fact, the use of BHT and BHA as food additives to neutralize or delay lipid peroxidation in the food industry has been shown to be associated with several serious health risks. Some epidemiological studies focusing on the medium and long-term health effects of BHT and BHA on animal models have proved that these products are characterized by high toxicity and carcinogenic or mutagenic effects (Lanigan & Yamarik 2002; Agyei et al. 2015).
These adverse health effects attributed to BHT and BHA and other commonly used synthetic antioxidants have put more pressure on nutraceutical, pharmaceutical, and food industries, and have made the need to identify new sources of natural antioxidants as safe alternatives even more appealing and urgent.
Regarding total polyphenols, very few studies have measured the polyphenol content in microalgae compared to other cellular constituents (Goiris et al. 2012). Our results showed TPP levels varying between undetectable values and a maximum of 8.9 mg EqGA/g DW (
Statistical analysis revealed that there was no significant correlation between antioxidant activity and polyphenol content in the crude extracts, either in terms of PPT (
The antibacterial activity assessment against several human pathogenic bacteria revealed a moderate antibacterial potential of the unicellular green alga
According to previous research published by Maadane et al. (2015), ethanolic extracts of
The antileishmanial activity of microalgal extracts was evaluated against promastigote forms of the parasites
At present, marine organisms are increasingly recognized by scientists as an interesting natural source of new bioactive products and a promising alternative in the therapy and control of leishmaniasis. However, very few reports have been published on the antileishmanial capacity of microalgal extracts (Pereira et al. 2015).
To the best of our knowledge, this is the first study to demonstrate the antileishmanial effect of the genus
In addition, the results revealed that most of the isolates tested exhibited a particularly targeted antileishmanial activity against
The present study provides important, clear and reliable information on 10 microalgal strains that colonize several Tunisian inland water bodies with respect to their potential for production of phycobiliproteins, and the evaluation of their antioxidant, antibacterial, and antileishmanial effects. The cyanobacterium
In view of the results obtained in the present work, we believe that the optimization of culture conditions, especially light and temperature, targeting the most potential strains will have an impact on increasing their potentiality. In addition, the use of genetic engineering as a measure to control the expression level of the genes encoding this activity would be of great benefit.