1 Introduction

Nanotechnology is an operation for the manufacture of materials ranged from 1 to 100 nm in size [1]. Nanoparticles display totally novel or enhanced characteristics, constructed on confident properties, like size, spreading, and morphology of the grains. Biological methods for nanoparticle synthesis using microorganisms [2,3,4], enzymes [5], and plant or plant extracts [6] have been suggested as possible eco-friendly alternatives to chemical and physical methods, which proved to be expensive and result in toxic byproducts [7]. They also have discrete physical, optical properties, and biochemical functionality due to diverse controlled size and shape [8].

Silver nanoparticles (AgNPs) have acquired increasing interest due to their unique physical properties, chemical reactivity, and potential applications as antimicrobial [9], anticancer [10], antioxidant [11], and antiviral [12]. The most widely known applications of AgNPs are in therapeutic manufacturing in medical device coatings, drug-delivery carriers, imaging probes, and implants of silver-impregnated polymers [13] to inhibit the occurrence of the infection. AgNPs are also the most effective in contradiction of bacteria, viruses, and other eukaryotic microorganisms among all metallic nanoparticles. This is mainly owing to the intrinsic inhibitory and bactericidal potential of silver, as well as due to their chemical stability, catalytic belongings, and dressed conductivity [14]. The antimicrobial activity of AgNPs synthesized via different biological approaches has been assessed against a wide range of multidrug-resistant bacterial and fungal pathogens. This includes Gram-negative bacteria such as Pseudomonas aeruginosa, Klebsiella pneumoniae, and Escherichia coli as well as Gram-positive bacteria such as Staphylococcus aureus [15]. In addition, AgNPs can also boost antifungal activity against numerous drug resistance fungal strains like Aspergillus fumigatus, Fusarium solani, Aspergillus niger, and Aspergillus flavus [16]. Presently, research tendencies are directed for screening and identifying new natural antioxidants from plant resources. Antioxidant activity in plant extracts is due to the redox potential of phytochemicals [6], which can play an important role in slaking singlet and triplet oxygen, disintegrating the peroxides, or neutralizing the free radicals. So, it is supposed that the favorite adsorption of an antioxidant component of a certain extract onto the surface of the nanoparticle is the main cause of the highest antioxidant activity of these nanoparticles [16]. The antioxidant activity of AgNPs proved by using many assays such as phosphomolybdenum assay, metal chelating activity, superoxide radical scavenging activity, DPPH radical scavenging activity, and ferric reducing/antioxidant power (FRAP) assays [17]. In a related context, AgNPs represent a novel class of antiviral agents due to their advantageous broad-spectrum antiviral activity [18, 19]; and because their antiviral action is occurring at non-cytotoxic concentrations compared to the current toxic broad-spectrum antivirals [20]. Singh et al. [14] mentioned a detailed review of the application of AgNPs as virucidal agents and demonstrated the effective inhibition of influenza viruses, which can be extended to other viruses as HIV, hepatitis C, and B viruses. The release of silver ions, cell membrane disruption, and DNA damage are the key mechanisms of AgNP action [18].

Basically, AgNP cytotoxicity is depending on their size and shape, capping or coating agent, concentration or dosage, surface charge, oxidation state, and the ability for agglomeration, as well as the pathogenic type for which AgNP toxicity is examined [21]. Numerous, in vitro, investigations have specified AgNP toxicity to mammalian cell lines, e.g., liver, lung, skin, and brain; the reproductive organs and vascular system cells as well [22]. However, a major concern of hazard practicing with respect to human health was prospected. In this context, a detailed review was conducted by Ferdous and Nemmar [23], who stated that AgNP toxicity and bio-distribution, in vivo, studies have proven Ag+ translocation and accumulation leading to toxicity for different organs. The same review emphasized that complete characterization of NPs and proper control of assays are mandatory when applying AgNPs on humans and the environment [23].

Cyanobacteria, previously known as blue-green microalgae, have a noteworthy attraction as a natural source of bioactive compounds with a comprehensive range of biological activities counting antimicrobial, antiviral, anticancer, antioxidant, and antiinflammatory possessions [3, 24, 25]. Cyanobacteria are rich in phycobiliproteins (PBPs), the unique photosynthetic pigments of cyanobacteria [26]. Spirulina platensis and Nostoc linckia are two common cyanobacterial species, and phycobiliproteins extracted from them displayed favorable health benefits for improving immune function and inhibiting the proliferation of cancer cells due to their fluorescent properties [26]. These pigments have been broadly used in the fields of foods, pharmaceuticals, and cosmetics as a natural alternative for synthetic colorants [27]. Owing to their photosensitive ability, phycobiliproteins have been proposed as a novel photosensitizer for photodynamic therapy [28, 29]; in the field of immunology, in addition to their role as an antioxidant and antiinflammatory agents [30]. In light of the above-mentioned literature, the present study is conducted to extract phycobiliproteins from S. platensis and N. linckia cyanobacterial species. The ability of these pigments to act as a biological reductant for AgNP biosynthesis is evaluated. Also, the antimicrobial, antioxidant, and antiviral properties of these AgNPs were assessed for potent biological applications.

2 Materials and Methods

2.1 Cultivation of Cyanobacterial Species

Axenic cyanobacterial cultures were obtained from the culture collection of Phycology Laboratory, Botany Department, Faculty of Science, Tanta University. Spirulina platensis (Gomont) was cultivated in a modified Zarrouk medium [31], while Nostoc linckia (Bornet) was cultivated in a BG11 medium [32]. For working cultures, 100 ml of stock inoculant corresponding to each species was cultured in 2-l Erlenmeyer flasks containing sterilized medium [33]. Cultures were then incubated under a surface light intensity of 45 μ mole photon m−2 s−1 at 30 °C and supplied with a mixture of 97% dry air and 3% CO2 to accelerate growth. The pumped air was sterilized first by passing through bacterial air filters of 0.45-μ pore diameter. The cultures were grown until the beginning of the stationary phase corresponding to each species. For this study, S. platensis biomass was harvested on the 16th day, and N. linckia biomass was collected on the 22nd day of growth, at which the maximum content of phycobiliproteins was achieved. Cyanobacterial biomass was harvested by centrifugation (Centurion Scientific, Model: CR2000, UK) at 2000g for 20 min. Pellet cells were rinsed, washed three times with distilled water, and re-suspended in sterilized distilled water to remove traces of growth medium [34].

2.2 Extraction of Phycobiliprotein Pigment

Crude phycobiliproteins from S. platensis and N. linckia were extracted according to Bennet and Bogorad [35]. Whereby, the freezing and thawing techniques of cell suspension (50 ml) in distilled water were manifested for two times at 24-h intervals to break the cells. The cells were then sonicated at a pulse of 60% for 15 min (Chromotech Ultra Sonic Processor). After sonication, the biomass was centrifuged at 2000g for 20 min to get the phycobiliprotein pigment. The procedures were repeated for 3 days for S. platensis and 7 days for N. linckia. Finally, the obtained volumes of the pigments were collected together and termed as “crude phycobiliprotein extract.” The pigment components were estimated spectrophotometrically at 615 nm for phycocyanin (PC), at 652 nm for allophycocyanin (APC), and 526 nm for phycoerythrin (PE). The following equations were applied to calculate the concentration of each component:

$$ \mathrm{PC}=\left({E}_{615}\hbox{---} 0.474\ {E}_{652}\right)/5.34 $$
$$ \mathrm{APC}=\left({E}_{652}\hbox{---} 0.208\ {E}_{615}\right)/5.09 $$
$$ \mathrm{PE}=\left({E}_{526}\hbox{---} 2.4\ \left(\mathrm{PC}\right)\hbox{---} 0.849\ \left(\mathrm{APC}\right)\right)/9.62 $$

where E is the absorbance value at each specific wavelength, and numbers are specific coefficients.

The crude phycobiliprotein extracts were then stored in a refrigerator at 4 °C in dark bottles until used for nanoparticle synthesis.

2.3 Biosynthesis of Silver Nanoparticles

For biosynthesis of AgNPs: of S. platensis and N. linckia crude phycobiliprotein extract (90 ml) was added to AgNO3 (10 ml). The reduction of silver metal ions (Ag+) to nanoparticles (Ag0) took place at room temperature and pH 7. The visual appearance of a yellowish to brownish color in the solutions was an evident indication of AgNP formation [36].

2.3.1 Characterization of AgNPs

Ultraviolet-Visual Spectroscopy

The developed color was monitored through aliquots sampling (0.2 ml) of the colloidal suspension diluted to 2 ml with deionized water and measured using ultraviolet-visual (UV-Vis) spectra of a double beam PC scanning spectrophotometer scanned at 200–800-nm range. The appearance of a plasmon resonance peak at 420 nm verifies the biosynthesis of AgNPs.

Fourier Transformer Infrared Analysis

Fourier transformer infrared (FTIR) spectrophotometer (AKX, Genesis Series Nicolet, IS-10 F) was used to identify the functional groups in S. platensis and N. linckia crude phycobiliprotein extract which were responsible for the reduction of silver ions. Along with the range of 500 to 4000 cm−1, the spectrum was chronicled.

Powder X-ray Diffraction Analysis

The X-ray diffraction (XRD) dimensions were performed (Philips-X’Pert MPD X-ray diffractometer) to determine the size and the crystalline shape of the AgNPs. The dried NP powder was coated onto a glass substrate while scanned in the region of 2θ, from 4 to 90° at 0.5°/min with the time constant of 2 s. The mean particle diameter of AgNPs was designed from the XRD pattern according to the line width of the maximum intensity reflection peak using Scherrer’s equation as:

$$ D=\left( K\lambda \right)/\left(\beta \mathrm{cor}\ \cos\ \uptheta \right) $$

“D” is the average crystal size, K is the Scherrer coefficient (0.89), λ is the X-ray wavelength (λ = 1.5406), °2θ is the Bragg’s angle, βcor is the corrected full width at half maximum (FWHM) in radians, with βcor = (β2sample − β2ref) 1/2, and β sample and β ref. are the FWHM of the sample and reference peaks, respectively.

Transmission Electron Microscopy

Transmission electron microscopy (TEM) analysis was employed to visualize the shape and confirm the size of the biosynthesized AgNPs. A drop of AgNPs on a carbon-coated TEM grid was allowed to dry at room temperature before analysis. TEM was carried out using JEOL-JEM 2100 Electron microscope functioning at 80 KV and a magnification of × 20,000 and a resolution up to 100 nm for micrographs.

Zeta Potential Analysis

Zeta potential of the biosynthesized AgNP aqueous suspensions was determined by placing 2 ml of the samples in a four-sided clear plastic cuvette in a Malvern zeta sizer instrument (Malvern instrumentation Co.) at 25 °C. The measurement was performed directly in the clear disposable zeta cell.

2.3.2 Applications of Biosynthesized AgNPs

Antimicrobial Activity of AgNPs

Five pre-identified bacterial pathogenic strains were obtained from Bacteriology Laboratory, Faculty of Science, Tanta University. Microbial suspensions were prepared in a sterile saline solution (0.9% w/v NaCl) at a turbidity of 106 to 108 CFU/ml. The Gram-negative bacterial strains Escherichia coli, Klebsiella pneumoniae, and Proteus vulgaris were cultivated in MacConkey solid medium, and Gram-positive bacterial strains Diplococci sp. and Staphylococcus aureus were grown onto nutrient agar medium [37]. One fungal strain, Candida albicans, was cultivated on Sabouraud Dextrose agar medium. The antimicrobial activities of the biosynthesized AgNPs were evaluated using the disk diffusion method [38]. Sterilized paper disks loaded with 15, 30, and 60 μg/disk of AgNPs were placed in the Petri dishes pre-inoculated with 100 μl of each microorganism suspension and then incubated at 37 °C. Positive control of standard antibiotics was tested and also negative controls of AgNO3 solution and cyanobacterial phycobiliprotein crude extract. After the incubation period, the inhibition zone diameters were measured and compared to the standard reference table of antibiotics [38]. The positive antibacterial controls were ciprofloxacin (CIP), doxycycline (DO), and ampicillin-sulbactam (SAM), while the antifungal positive controls were fluconazole (FLU), itraconazole (ITC), and metronidazole (MT).

2.3.3 Antioxidant Activity of AgNPs

DPPH Radical Scavenging Activity

The capacity of AgNPs to reduce 2,2-diphenyl- picrylhydrazyl (DPPH) stable free radical was assessed as described by Blois [39]. Three hundred microliters of AgNPs at various concentrations was mixed with 2.7 ml of DPPH reagent solution and allowed to stand for 30 min in the dark at room temperature. The absorbance of the samples was measured at 517 nm, and DPPH only was also detected as a negative control. The antioxidant capacities of the samples were compared with ascorbic acid as a standard. The IC50 value of DPPH and ascorbic acid was also calculated. The DPPH scavenging activity was calculated as a percentage according to the following formula:

$$ \mathrm{Inhibition}\ \mathrm{of}\ \mathrm{the}\ \mathrm{DPPH}\ \mathrm{activity}\ \left(\%\right)=\left({A}_{\mathrm{c}}-{A}_{\mathrm{s}}\right)/\left({A}_{\mathrm{c}}\right)\times 100 $$

where Ac is the absorbance of the control and As is the absorbance of the sample.

Total Antioxidant Capacity Assay

The total antioxidant activity of AgNPs was evaluated by the phosphomolybdenum method [40]. Different concentrations of AgNPs were prepared by dissolving a known weight of AgNPs in 10 ml of 1 mM dimethyl sulfoxide. Three hundred microliters of each concentration was added to 3 ml of phosphomolybdate reagent solution (0.6 M sulfuric acid, 28 mM sodium phosphate, and 4 mM ammonium molybdate). The reaction was protected from light, and then the vials were incubated for 90 min in a water bath at 95 °C. The absorbance was measured at 695 nm against a blank after cooling the mixture at room temperature. Ascorbic acid was used as a reference. Results were expressed as mg ascorbic acid/g dry wt of the NPs (mg AAE/g DW).

The Ferric Reducing Antioxidant Power Assay

The chelation activity of ferrous ions by AgNPs was estimated as described by Do et al. [41]. First, 1 ml each of different AgNP concentrations was added to 2.5 ml of sodium phosphate buffer (0.2 mM, pH = 6.6) and 2.5 ml of potassium ferrocyanate (1%) then the mixture was incubated for 20 min in a water bath at 50 °C. After that, 2.5 ml of tricholoroacetic acid (10%) was added before centrifugation of the mixture solution at 500 g for 10 min. Then, 5 ml of the supernatant was mixed with 5 ml dist. H2O and 1 ml of ferric chloride solution (0.1%). The absorbance of the mixture was measured at 700 nm. Higher absorbance values indicate increased reducing power. The results were reported as ascorbic acid equivalents (AAE) per gram dried sample (mg AAE/g DW).

2.3.4 Antiviral Activity of AgNPs

Mammalian Cell Culture Conditions

Mammalian cells, hepatocellular carcinoma, and HepG2 were obtained from VACSERA, Egypt. And then were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 2 mM of L-glutamine, 10% of fetal bovine serum (FBS), and 1% penicillin-streptomycin solution in 50-ml cell culture flasks, incubated at 35 °C in a humidified atmosphere of 5% CO2 and 95% air for 24 h. For experimental purposes, cells were cultured in 6-, 12-, or 96-well plates.

2.3.5 Cytotoxicity Test

Through this test, the maximum non-cytotoxic concentration of the test sample was determined. This concentration was used for all tests without having any toxic effect on the cells [42]. Staining vital cells with neutral red has been known for a long time. Only vital cells can incorporate neutral red into the liposome, so, the amount of absorbed stain is proportional to the number of vital cells [43]. Peripheral Blood Mononuclear Cells (PBMC) were plated into 96-well plates at a cell count of 5 × 104 cell ml−1. After 24 h of incubation, the cells were semi-confluent, and serial dilution of the AgNPs in culture medium was added. The treated cells were then incubated for 3 days at 37 °C and 5% CO2 supply, during which they were exposed to a serial dilution of the AgNPs. After that, 100 μl of neutral red was added to every well, and the plates were incubated for 3 h. Thereafter, cells were fixed with100 μl fixative solution (0.5 formaldehyde and 1% CaCl2) for 1 min. Then, each well was supplemented with 100 μl solution of ethanol with 1% acetic acid while shaking for 5 min. Finally, the adsorption of plates was measured at 450 nm. Plotting concentrations against the percentage of cell survival gave a cytotoxicity curve, from which the maximum non-cytotoxic concentration was determined and used afterward in the following experiments. From the same curve, the IC50 toxic concentration value was also calculated [43].

Determination of Virus Concentration

Hepatitis C virus (HCV) concentration was firstly determined by real-time polymerase chain reaction (RT-PCR).

2.3.6 Antiviral Assay

Non-toxic concentrations of the AgNPs in 50 ml of DMEM-supplemented medium were added simultaneously with the virus (infected serum, 8.3 × 106 copies/ml, genotype 4a) to the host cells (HepG2, 105 cells/ml). After 90-min incubation period, the infected cells were washed three times with 1 ml of phosphate-buffered saline (PBS) and further cultured for 3 days at 37 °C and 5% CO2, and then total RNA extraction had done.

Extraction of RNA from HepG2 Cells

The positive and/or negative HCV RNA strand was separated from HepG2 cells, which was reported by Lohr et al. [44]. Briefly, cells were precipitated and washed in the same buffer to remove adherent viral articles, then lysed in 4 mol/l guanidinium isothiocyanate (containing 25 mmol/l sodium citrate, 8 μl β-mercaptoethanol, 5% sacrosyl, and 100 μl sodium acetate). Five hundred microliters of phenol (pH = 4) and 200 μl of chloroform were added then vortexed. The lysed cells were centrifuged at 12000 rpm for 20 min at 4 °C. The upper aqueous layer was transferred to new falcons and mixed with the same volume of isopropanol and incubated overnight at 20 °C. RNA was precipitated by centrifugation at 12000 rpm for 20 min at 4 °C. The precipitate was washed with 70% ethanol, and the previous centrifugation procedures were repeated twice.

PCR of Genomic and Antigenomic RNA Strands of HCV

Reverse transcription polymerase chain reaction (RT-PCR) was done following the method of Lohr et al. [44] with some modification. Reverse transcription was performed in 25-μl reaction mixture which contains 20 U of AMV (alfalfa mosaic virus) reverse transcriptase, 2 mmol/l from each of dNTP (Deoxy Nucleotide Triphosphate; Promega, Madison, WI, USA) with 50 pmol of reverse primer 1CH used as plus strand and 50 pmol of forwarding primer 2CH used as a minus strand. This reaction had been incubated for 60 min at 42 °C then denatured for 10 min at 98 °C. Two rounds of PCR with two pairs of nested primers were used for amplification of highly conserved 5′–UTR sequences. A reaction mixture of 25 μl that contains 50 pmol from each of 2CH forward primer and P2 reverse primer was carried out for first-round amplification. 0.2 mmol/l of dNTP, 10 μl from RT reaction mixture (template and 2 U of Taq DNA polymerase in 1X buffer) were added to the reaction mixture. Then a protocol of 1 min at each of 94 °C, 55 °C, and 72 °C for 30 cycles, which was described as a thermal cycling protocol, was applied. Then the second round amplification has been carried out as the first one, excluding the use of nested reverse primer D2 and the forward primer F2 at 50 pmol. A fragment of 174 bp had been recognized in positive samples. The primer sequences used in the reaction were:

  • 1CH: 5′-GGTGCACGGTCTACGAGACCTC-3′,

  • 2CH: 5′-AACTACTGTCTTCACGCAGAA-3′,

  • P2: 5′-TGCTCATGGTGCACGGTCTA-3′,

  • D2: 5′-ACTCGGCTAGCAGTCTCGCG-3′ and

  • F2: 5′-GTGCAGCCTCCAGGACCC-3′.

Strand-Specific RT-PCR

The final PCR product was checked by the RT-PCR, which constructed on SYBR Green 1 dye and light Cycler fluorimeter method that use standard serum samples infected by HCV. During PCR of the 5′ HCV non-coding (NC) region, the SYBR Green 1 dye binding to double-stranded DNA for the synthesis of amplicon that continuously monitored and specifically verified by temperatures of amplicon melting. A serial of a modified synthetic HCV 5′ NC RNA of 10-fold dilutions was created with an external standard curve. After the RT step, the first-round PCR, which uses 2CH and P2 primers in 25-μl reaction mixture that contains 3.5 μl cDNA, 50 pmol of 2CH forward primer, 50 pmol of P2 reverse primer, and 12.5 μl of SYBR Green master mix, was done, then the previous thermal cycling protocol of 30 cycles of 1 min at 94 °C, 55 °C and 72 °C with final extension made for 5 min at 72 °C [45].

Statistical Analysis

One way analysis of variance (ANOVA) was performed for the results of the antimicrobial test. The difference between means pigments and antioxidant results (total antioxidant capacity (TAC) and FRAB) in both cyanobacteria series was tested by using an unpaired t - test. All results were expressed as a mean of three replicates ± standard deviation (SD) and statistically evaluated using SPSS 16.0 software SPSS (2006) [46].

3 Results

3.1 Estimation of Pigment Concentration in the Crude Phycobiliprotein Extract

The PC, APC, and PE pigment components in the crude phycobiliprotein extract are shown in Table 1 for S. platensis and N. linckia. N. linckia exhibited a highly significant total pigments composition (0.307 mg/ml), of which PC value was the maximum (0.158 mg/ml).

Table 1 Concentration of pigments (mg/ml) in the crude phycobiliprotein extract of S. platensis and N. linckia
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3.2 Visual Biosynthesis and Ultraviolet-Visual Spectroscopy of AgNPs

Reduction of the silver ion to AgNPs by crude phycobiliprotein extracts of S. platensis and N. linckia was followed by a visual change of solution color to reddish-brown, as shown in Fig. 1. The change in color indicated the formation of AgNP solution through the biotransformation of the ionic silver (Fig. 1). For the characterization of the formed AgNPs, the UV-Vis absorption peak was monitored at a time interval of 2, 6, 12, and 24 h for S. platensis AgNPs (Fig. 2a) while at 24, 48, 72, and 96 h for N. linckia ones (Fig. 2b). The surface plasmon resonance bands at 420 nm indicated the production of AgNPs of S. platensis and N. linckia phycobiliprotein crude extracts. Although the peak intensity of both species was nearly the same, the formation of AgNPs by S. platensis phycobiliproteins was markedly faster. By increasing the period of incubation time, the peak at 420 nm got sharper and narrower because of the completed construction of AgNPs. These results also indicated that AgNPs were dispersed in the aqueous solution with no evidence for aggregation, and its formation was dependent on the time and the algal extract source.

Fig. 1
figure 1

S. platensis crude phycobiliprotein extract (a) and its biosynthesized AgNPs (b); N. linckia crude phycobiliprotein extract (c) and its biosynthesized AgNPs (d)

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Fig. 2
figure 2

UV-Vis spectra of AgNPs synthesized by crude phycobiliprotien extract of S. platensis (a) and N. linckia (b)

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3.3 Fourier Transformer Infrared

The functional groups of the biomolecules in S. platensis and N. linckia phycobiliprotein extracts were identified by FTIR. The FT-IR spectrum of S. platensis AgNPs (Fig. 3a) showed various absorption peaks (cm−1) at 3796 and 3430 which indicates the presence of OH strong stretching band of alcohols or phenols, respectively. The peak at 2925 represents N–H group (amino acids), and at 2426 and 2360 represent C=O of the carboxylic anion. The peak at 1634 indicates the presence of broad stretching aliphatic bond of carboxylate group C=O; at 1383 represents a strong sharp band of C–H or CH3 methyl group; at 1153 is a broadband carboxyl of carboxylic acid C–O–C and at 875 is a broad weak band of C–O–S sulfate group. Similarly, the FT-IR spectrum of N. linckia AgNPs (Fig. 3b) showed strong absorption peaks at 3872, 3384, and 3210 cm−1 due to O–H stretching vibration presence of alcohols or phenols, while the peaks at 2964 and 2924 can be assigned to stretching broadband of N–H group (amino acids). The peaks at 2426, 2363, and 1641 cm−1 represented the protein amide, mainly C=O stretching broad, which may be due to the N–H bending vibration of the carbonyl β unsaturated ketone amide and secondary amines (protein, lipid). The peak at 1383 represents a strong, sharp band of C–H or CH3 methyl group, and the bands at 1044 and 877 were allocated to the stretching vibrations of primary and secondary amines, respectively.

Fig. 3
figure 3

FT-IR spectra of AgNPs synthesized by phycobiliprotein crude extract of S. platensis (a) and N. linckia (b)

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3.4 Powder X-ray Diffraction

The XRD analysis showed intense peaks corresponding to 111, 200, 220, and 311 Bragg reflections of the biosynthesized AgNPs (Fig. 4a and b), which indicated a crystalline nature of the formed nanoparticles. The average crystal size was calculated from the FWHMs of the diffraction peaks using Scherrer’s equation. The FWHMs of all detected peaks of S. platensis AgNPs were at 38.6° (111), 44.8° (200), 64.8° (220), and 77.9° (311), while the FWHMs of all peaks of N. linckia AgNPs were at 38.55° (111), 44.6° (200), 64.5° (220), and 77.3° (311) in °2θ which were used for size evaluation. The average estimated size was 21.05 nm and was 20.84 nm for S. platensis and N. linckia AgNPs, respectively. A more broadening of the Bragg peaks indicated more reduction in the grain size.

Fig. 4
figure 4

XRD analysis of AgNPs synthesized by crude phycobiliprotein extract of S. platensis (a) and N. linckia (b)

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3.5 Transmission Electron Microscopy

TEM is a commonly used technique for characterizing the size and shape of the biosynthesized nanoparticles. All the particles were spherical. The particle size distribution of S. platensis AgNPs was ranged from 15.1 to 27.4 nm with a mean particle size of 21.211 nm. For the AgNPs formed by N. linckia, the particle size distribution was in the range of 16.3 to 25.8 nm with a mean particle size of 21.052 nm (Fig. 5a and b).

Fig. 5
figure 5

TEM image of AgNPs by phycobiliprotein crude extracts of S. platensis (a) and N. linckia (b)

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3.6 Zeta Potential Analysis of AgNPs

Zeta potential analysis verified the stability of the biosynthesized AgNPs. A net charge of − 15.902 ± 4.87 mV and − 16.811 ± 6.99 mV with a conductivity of 0.225 and 0.23 mS/cm were detected for S. platensis and N. linckia AgNPs, respectively (Fig. 6a and b). The zeta potential of nanoparticles that is greater than + 30 mV or less than − 30 mV has sufficient electrostatic repulsion to remain stable in solution [25].

Fig. 6
figure 6

Zeta potential analysis of AgNPs synthesized by crude phycobiliprotein extracts of S. platensis (a) and N. linckia (b)

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3.6.1 Antimicrobial Activity of AgNP

The bactericidal activity of S. platensis and N. linckia AgNPs was performed against five pathogenic bacterial species: two were G+ve bacteria of Diplococci sp. and Staphylococcus aureus and three were G–ve bacteria of E. coli, K. pneumoniae, and P. vulgaris in addition to Candida albicans fungal strain. By increasing the concentration of AgNPs, the diameter of the inhibition zones also increased (Table 2). At the lowest AgNP concentration of 15 μg/ml, the diameter of different inhibition zones was insignificant compared to those recorded for the negative controls, especially with S. platensis AgNPs. At the highest concentration of 60 μg/ml, an elevated inhibitory effect was observed for AgNPs biosynthesized by S. platensis crude phycobiliprotein extract against P. vulgaris (15.5 mm), Diplococci sp. (14 mm), and S. aureus (12.1 mm). At the same concentration of N. linckia AgNPs, a higher significant inhibition zones were recorded against Diplococci sp. (20 mm), S .aureus (17.8 mm), P. vulgaris (15 mm), and K. pneumonaie (13.75 mm) compared to the antibacterial positive controls. A moderate antifungal activity against C. albicans was recorded for AgNPs of N. linckia (10.5 mm) and S. platensis (9.8 mm) compared to the significant antifungal controls, especially for MT and FLU. As a negative control, AgNO3 solution and crude phycobiliprotein extracts of S. platensis and N. linckia treatments showed insignificant, weak inhibitory effects against G−ve rather than G+ve bacterial as well as the fungal strain tested pathogens (Table 2).

Table 2 Diameters of inhibition zones (mm) of AgNPs synthesized by crude phycobiliprotein extracts of S. platensis and N. linckia compared to antibiotics, antifungal, and negative controls
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3.6.2 Antioxidant Activity of AgNPs

DPPH Radical Scavenging Activity

The IC50 value of DPPH scavenging activity was considerably higher (133.8 μg/ml) for S. platensis phycobiliprotein AgNPs than that of AgNPs synthesized by N. linckia crude phycobiliprotein extract, which records 77.3 μg/ml. In the present study, increased concentrations of AgNPs showed comparatively higher DPPH antioxidant activity (Table 3) compared to ascorbic acid (AA) as standard antioxidant. The IC50 value of ascorbic acid was 67 μg/ml.

Table 3 DPPH radical scavenging activity of AgNPs synthesized by crude phycobiliprotein extracts of S. platensis and N. linckia
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Total Antioxidant Capacity

By increasing AgNP concentration, the total antioxidant capacity increased too (Fig. 7a). The biosynthesized AgNPs showed significant phosphomolybdenum reduction activity (at p ≤ 0.001), recording 44.57 and 45.13 mg AAE/g DW for S. platensis and N. linckia AgNP crude phycobiliprotien extracts, respectively at 300 μg/ml.

Fig. 7
figure 7

Antioxidant activity of AgNPs synthesized by crude phycobiliprotein extracts of S. platensis and N. linckia: Total antioxidant capacity (a), ferric reducing antioxidant power (b). All mean values are of significant difference at p ≤ 0.001

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Ferric Reducing Antioxidant Power

Similar behavior was observed using FRAP assay as that observed for TAC assay where the antioxidant activity for both S. platensis and N. linckia phycobiliprotein AgNPs exhibited significant difference values of 34.75 and 35.13 mg AAE/gDW, respectively at the concentration of 300 μg/ml (Fig. 7b).

3.6.3 Antiviral Activity of AgNPs

Cytotoxicity of the Biosynthesized AgNPs on HepG2 Cell Line

The cytotoxicity of N. linckia and S. platensis phycobiliprotein extract AgNPs on human liver carcinoma (HepG2) cell line was assessed using the neutral red assay. At lower concentration of 16 μg/ml, the cell viability recorded 90% ± 0.2 and 93% ± 0.2; while at 1000 μg/ml, the cell proliferation decreased to 48.1% ± 0.5 and 53.3% ± 0.4 for S. platensis and N. linckia AgNPs, respectively. As the AgNP concentration increased, the cell viability decreased in a direct dose-response relationship (Fig. 8). The concentration necessary to produce 50% of tumor cell death (IC50) was 992 μg/ml for S. platensis AgNPs, while the same concentration for N. linckia AgNPs was not detected indicating that N. linckia AgNPs were safer in the tested concentration range. Thus, the concentration of 16 μg/ml AgNPs was chosen for the rest of the experiment as the safe dose (Fig. 8).

Fig. 8
figure 8

Cytotoxicity of AgNPs biosynthesized by S. platensis and N. linckia phycobiliprotein crude extract on human liver cancerous (HepG2) cell line

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Antiviral Activity Assay Using RT-PCR Technique

Results indicated that the antiviral activity of Ribavirin, as a standard antiHCV drug, recorded 66.673%. AgNP phycobiliprotein crude extract of N. linckia recorded 64.976% which compared effectively to Ribavirin (at p ≤ 0.05), while S. platensis AgNPs showed a lower potent activity of 48.334% (Table 4).

Table 4 Antiviral assay of AgNPs synthesized by crude phycobiliprotein extracts of S. platensis and N. linckia on hepatitis C virus
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4 Discussion

In cyanobacteria, phycobiliproteins are responsible for harvesting the light and deliver it to chlorophyll. Phycobiliprotein pigments (PBPs) can be classified into three types of phycocyanin (PC), phycoerythrin (PE), and allophycocyanin (APC), according to their maximum absorbance [27]. Many researchers reported the synthesis of AgNPs by cyanobacterial filtrate and biomass [47, 48]; only a few reports are available regarding the phycobiliproteins of cyanobacteria for the synthesis of NPs [15]. The synthesis of AgNPs was originally determined by the visual change in the color of silver nitrate solution from colorless to reddish-brown. Our result clearly showed that phycobiliprotein crude extract from N. linckia and S. platensis efficiently reacted with AgNO3 to form AgNPs by a reduction process in a range of 24 to 96 h of incubation time. By using the UV-Vis spectroscopy, the surface plasmon band of the biosynthesized AgNPs was at 420 nm throughout the reaction period, indicating that NPs were stable and well dispersed in the solution. Electrostatic interaction between the silver ions and the negatively charged carboxylate groups, which are present in phycobiliproteins, is thought to be the first step that involves the trapping of metal ions on the surface of algal cells [49]. The spectra of AgNPs were also dependent on the size, shape, and incubation time of the particles. The results showed that reduction of Ag+ ions into Ag0 started within 2 h after the addition of AgNO3 solution to S. platensis phycobiliprotein extract and completed after 24 h of incubation, while started within 24 h for N. linckia phycobiliprotein extract and completed after 96 h (Fig. 1a and b). This indicates the rapid synthesis and stability of the biosynthesized AgNPs. The obtained results were compatible with those reported in [15, 50].

FTIR indicated that AgNPs biosynthesized by S. platensis and N. linckia phycobiliproteins have nearly the same peaks, which represent similar functional groups, but most important changes in stretching frequencies were observed after the synthesis of AgNPs. The involvement of several functional groups such as hydroxyl, carboxyl, and carbonyl groups of amino acids and proteins were logged that might be responsible for the reduction of silver ions and also for capping of the bio-reduced AgNPs [51] as well as stabilization of these NPs for long duration by avoiding their aggregation and growth in the solution. The presence of some aromatic amino acids like tryptophan, phenylalanine, and tyrosine in the composition of phycobiliproteins and which induces the formation of AgNPs makes phycobiliproteins play a great important role in the synthesis and stabilization of metal NPs [52]. In agreement with our results, Mubarak-Ali et al. [48] reported the existence of cyanobacterial protein covering around Cd–S nanoparticles from cyanobacteria Oscillatoria welli, which was accounted for the NP biosynthesis. These protein molecules boosted the formation of AgNPs with a particularly small size and narrow particle distribution. On the other side, the presence of sulfonic and hydroxyl groups is useful for the biosynthesis of AgNPs with rather larger particle size [48]. In addition, besides providing stability and capping, the presence of phycobiliprotein biomolecules may offer anchoring ability to the developed NPs on bacterial cell membranes, enabling them to achieve antibacterial activity as reported by El-Naggar et al. [15].

In this study, the average size of the developed AgNPs was 21.05 and 20.84 nm for S. platensis and N. linckia respectively, as estimated from the FWHM of the four peak planes of silver ions. The crystalline nature of the nanoparticles by having a face-centered cubic (FCC) structure was confirmed by XRD analysis. AgNPs were found to be spherical in shape and well distributed in the solution. These results were in conformity with those reported by [53, 54]. TEM technique was also applied in this study to assess the particle size measure and the morphological shape of the biosynthesized AgNPs. From TEM images, AgNPs were spherical and well distributed with an average size below 25 nm for AgNPs of both S. platensis and N. linckia extracts. These results were in agreement with those reported by Shivaji et al. [53], who assessed that AgNPs are predominantly spherical.

The zeta potential is a physical property that is exhibited by any particle in suspension and predicting the long-term stability of these particles. The surface potential of the particles is reflected by its zeta potential and is influenced by the interface changes with the dispersing medium, due to the dissociation of the functional groups on the particle surface or to the adsorption of ionic species present in the aqueous dispersion medium [55]. A relatively high value of the zeta potential is important for good physical and chemical stability of the colloidal suspension because large repulsive forces tend to prevent aggregation due to incidental collisions of adjacent nanoparticles. Nanoparticles with zeta potential signal greater than + 30 mV or less than − 30 mV are considered strongly cationic and strongly anionic, respectively [15, 25]. In this connection, the biosynthesized AgNPs by S. platensis and N. linckia phycobiliproteins are considered strongly anionic which is important in the biological application, according to Aiad et al. [56].

The antimicrobial activity of biologically fabricated AgNPs has been reported by many researchers [54, 57]. As the extensive use of several synthetic antimicrobial agents has led to multidrug resistance in a number of bacterial strains, the results of the present study seems to be remarkable because the synthesized AgNPs showed strong antibacterial activity against five multidrug-resistant bacteria, E. coli, K. pneumoniae, P. vulgaris, S. aureus, and Diplococci sp., especially for N. linckia at the highest tested concentration of 60 μg/disk. Generally, as the concentrations of AgNPs increased, the diameter of the inhibition zone was increased too. The recorded variances in sensitivity among the tested bacteria against AgNPs might be due to their differences in the building of their bacterial cell wall. Gram-positive bacteria have a thick peptidoglycan layer (20–30 nm) between the cytoplasmic membrane and the outer—non-lipid—membrane, while Gram-negative bacteria possess a thin peptidoglycan layer of about 3 nm in thickness and have an outer lipid membrane. This may be the cause of the high antibacterial activity of AgNPs on S. aureus and Diplococci sp. G+ve strains. For such strains, this effect was significantly pronounced even at the lowest tested concentration of 15 μg/disk compared to the negative controls (especially for N. linckia AgNPs, implying the antimicrobial efficacy of phycobiliproteins as nanoform). The small size of the volume area of AgNPs and its crystalline structure are responsible for their antimicrobial potentials and can change the membrane permeability by causing perforations/pits in the bacterial cell wall, and thus releasing vital membrane proteins and lipopolysaccharide molecules [47]. Other scientists suggested that the cell damage may be caused by the interaction of the released silver ions, Ag+, with phosphorus and sulfur-containing compounds such as DNA and proteins, which in sequence lead to the detention of cell metabolic activities [14, 15]. Another potential pathway of the antimicrobial action is that, upon contact of AgNPs with bacteria, Ag+ ion release, disturb the reduction of intracellular O2, and interact with respiratory chain proteins on cell membranes, thus trigger free radicals and reactive oxygen species (ROS). This induced cellular oxidative stress could modulate the intracellular signaling routes of microbes’ cells in the direction of apoptosis [58]. All these findings supported the potential antibacterial activity of AgNPs synthesized by phycobiliprotein extract from S. platensis and N. linckia.

In the present study, the DPPH scavenging method confirmed that AgNPs have antioxidant activities due to the presence of functional groups on their surface [59]. DPPH is a stable nitrogen-centered radical; the higher the antioxidant activity of the sample is achieved by its higher DPPH radical scavenging activity. This nominated AgNPs of N. linckia phycobiliproteins to be a more active antioxidant than S. platensis ones. Nevertheless, both NPs presented potent significant antioxidant capacity by using phosphomolybdenum and FRAP assays. It should be noted that in the ferrozine and phosphomolybdenum assays, the electron transfer occurs at different redox potentials, and in each case, the reducing activity is governed by the structure of the antioxidant (AgNPs) under study [60].

AgNPs of S. platensis was more effective on HepG2 cell line than AgNPs of N. linckia recording a cell viability percentage of 93% at 16 μg/ml concentration of AgNPs. The inhibitory effect was more pronounced at higher concentrations of AgNPs, leading to nearly complete mortality of the cells. This indicated that AgNP cytotoxicity on HepG2 is mainly mediated by the release of silver ions (Ag+) from the particle surface and their consequent adsorption to the surfaces of the cells as a source of toxic Ag+ ions, or, in the presence of oxygen, ligands, or organisms; these toxic Ag+ ions are formed upon oxidative dissolution [61].

The AgNPs biosynthesized by S. platensis and N. linckia crude phycobiliprotein extracts attained a dose-dependent inhibition on the hepatitis C virus. The AgNPs affected viral penetration at 16 μg/ml (the pre-determined safe concentration) when added to the cells during in vitro infection, as a co-treated method [62]. N. linckia AgNPs recorded a higher antiviral percentage of 64.976% compared to Ribavirin (standard drug for HCV). This may be due to the smaller particle size of N. linckia (20.48 nm) and its lower zeta potential value (− 16.8 ± 6.99 mV) compared to S. platensis AgNPs. These features enabled their augmentation and penetration through the virus molecules and triggered a more efficient inhibition of the viral infection activity. As it has been established, smaller AgNPs ensures a faster rate of Ag+ distribution into the around micro-environment compared with larger sized NPs [63]. Their large surface area to volume ratio enables their higher bioavailability, augmented distribution, and consequently, their cytotoxic activity [63]. In agreement with our results, Sun et al. [64] reported that AgNPs could inhibit the replication of HIV and display promising cytoprotective activities toward HIV-infected T cells. Also, they found that AgNPs could bind to the viral particles of HIV with a regular spatial arrangement and were able to prevent the virus from binding to the host cells. El-Sheekh et al. [65] investigated both Ag2O/AgO-NPs and Au-NPs from two blue-green algal strains, Oscillatoria sp. and Spirulina platensis, against herpes simplex virus (HSV-1). Results indicated that 90% reduction value in the cytopathic effect (CPE) of HSV-1 was achieved when applying both types of nanoparticles at 31.25-μl dose concentration, with a reduced rate of 49.23% for Ag2O/AgO-NPs and 42.75% for Au-NPs, respectively.

5 Conclusion

The present study verified that S. platensis and N. linckia crude phycobiliprotein extract can biosynthesize AgNPs with small and stable properties. The obtained results displayed the potential antimicrobial and antioxidant capacity of these AgNPs. Moreover, the biosynthesize AgNPs exhibited potent antiviral activity against HCV and a strong cytotoxicity effect on HepG2 cell lines. The study recommended using cyanobacterial cells as commonly biological resources in the greenfield of nanotechnology for multi-purpose medicinal and pharmaceutical applications. Future studies are still in need to declare the intact cellular uptake mechanism(s) by which AgNPs exert their biological antiviral action and also to assess their safety limits for treatment, especially with human contact.