Reversible encapsulation of silver nanoparticles into the helix of amylose (water soluble starch)

Abstract

Natural biodegradable polymeric starch capped Ag-nanoparticles (AgNPs) were prepared by using extract of Dioscorea deltoidea in the presence of starch. UV-visible spectroscopy, transmission electron microscopy (TEM), X-ray diffraction (XRD), energy dispersive X-ray spectroscopy (EDX), scanning electron microscopy (SEM), Fourier-transform infrared (FT-IR) spectroscopy, and digital images were used to determine the morphology and chemical composition of the as prepared AgNPs. The kinetics and morphology of the nano-materials (spherical, rod, triangular, irregular, truncated triangular, hexahedral, mono-dispersed, and aggregated) were discussed in terms of the [extract], [Ag⁺], and [starch]. Iodometric titration was used to confirm the reversible encapsulation of the AgNPs inside the helical structure of amylose. TEM images also suggest that the morphology of the encapsulated AgNPs entirely changes in comparison with the non encapsulated AgNPs. The starch functionalized AgNPs could be used for drug delivery, with the nucleation and aggregation controlled through fusogenic behaviour.

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1 Introduction

The use of different stabilizers and/or capping agents (aqueous extracts of plants, surfactants, proteins, lipids, synthetic and natural polymers) in the synthesis of advanced noble metal nanomaterials has been the subject of various investigations from the last two decades.^1–5 The characteristics of each stabilizer have a significant impact on the shape, size, aggregation, and the size distribution of the synthesized materials.^6–10 For the synthesis of AgNPs, environmentally benign biodegradable non toxic natural polymers, especially polysaccharide, have an edge over other stabilizers due to the weak interactions with the nanoparticles.^11–13 Raveendran et al. in their pioneering work suggested that the polysaccharides containing large numbers of –OH groups, formed inter and intra chain hydrogen bonds and acted as a template for nanoparticle growth.^14,15 It is well known that carbohydrates are the most extensively used natural biopolymer for the synthesis of silver polymeric nanocomposites.^16–20 Starch is a biodegradable natural polymer of α-D-glucose. It can be separated into two fractions (amylose, 10–20%, which forms a colloidal dispersion in hot water and amylopectin, 80–90%, completely water insoluble). It is well known that amylose in starch is responsible for the formation of a deep blue color in the presence of iodine–potassium iodide reagents. Amylose (the water soluble constituent of starch) holds great promise as a template for AgNPs because of its low toxicity and highly regular structure allowing it to host AgNPs and form stable nanocomposites. The nanomaterials either bind to the surface as nanospheres or are encapsulated inside as nanocapsules in the helix of amylose.^21–23 On the other hand, starch also forms an intensely blue colored starch–iodine complex with iodine in water. The starch forces and/or encapsulates the iodine atoms into a linear arrangement in the central groove of the β-amylose coil, which is a soluble starch. The blue complex is unstable in an excess of reducing agent and has been used as an indicator in redox titrations.

Zboril et al. pointed out that polymeric materials exhibit different features due to the hydroxyl groups of the organic network as compared to the monomers (the polymer polyacrylamide is completely safe for humans, whereas the acrylamide monomer is toxic).²⁰ Vimala and her coworkers used cross-linked poly(acrylamide) from different carbohydrates (gum acacia, carboxymethylcellulose and starch) with N,N-methylenebisacrylamide for the synthesis of AgNPs as a stabilizer. They suggested that the amide and hydroxyl groups of the cross-linked polymers enhanced the stability and antimicrobial properties of the as prepared AgNPs inside the matrix.²⁴ The silver/biocompatible biodegradable polymeric nanocomposites (dextrose and poly(γ-glutamic acid),²⁵ poly(ethylene glycol dimethacrylate),²⁶ chitosan²⁷ and others²⁸) are excellent vehicles for incorporating a wide variety of additives, antioxidants, antifungal agents, antimicrobials, and drugs which are important in food preservation and drug delivery systems.

Our aim is to avoid the use of toxic stabilizer(s) in the synthesis of water soluble and biocompatible Ag-nanomaterials that can be used in biological systems.^29,30 Now, we use natural biodegradable water soluble starch (biocompatible with humans) as a stabilizer and/or capping agent. In this paper we report a simple chemical reduction method for the synthesis of AgNPs by using the aqueous extract of Dioscorea deltoidea leaves, and starch as the reducing and stabilizing agents. This paper also describes the reversible encapsulation of AgNPs into the helix of amylose. Starch capped nanomaterials could be used as models for appropriate biomedical applications in nanotechnology.

2 Experimental

2.1 Materials and methods

All glassware was washed with aqua regia (3 : 1 HCl and HNO₃) and rinsed with water (double-distilled, CO₂ free and deionised) prior to use. Silver nitrate (AgNO₃, BDH, 99.99%) and starch (stabilizer) were used as received without any further purification. A starch solution (2.0% w/v) was prepared by dissolving the required amount in deionized water and heating for 15 min in a 500 cm³ Erlenmeyer flask with constant rapid stirring to avoid the aggregation of starch. Dioscorea deltoidea tubers were collected from the Poonch district of Jammu and Kashmir, India. Approximately 10 g of Dioscorea deltoidea tuber was washed with water, chopped into fine pieces then soaked in 500 cm³ distilled water, and heated for 20 min in a water bath at 60 °C. The solution was cooled, decanted and filtered through Whatman no. 1 filter paper. The filtrate was stored in an amber colored airtight bottle to avoid the intervention of photochemical reactions.

2.2 Preparation and characterization of AgNPs

Preliminary observations showed that the extract plays an important role in the preparation of different colour silver sols with and without starch. In a typical experiment, extract (33.3% v/v) was added into a reaction mixture containing AgNO₃ (5.0 cm³ of 0.01 mol dm⁻³) + water (required volume for dilution) + starch (if needed). Surprisingly, the colourless reaction mixture became a transparent yellowish brown both in the absence and presence of starch indicating that the reduction of Ag⁺ ions into Ag⁰ leads to the formation of AgNPs.^30,31 Transmission electron microscopy (TECHNAI-320 KV JAPAN, operating at 80 kV system equipped with energy dispersion X-ray spectroscopy), a Rigaku X-ray diffractometer operating at 40 kV and 150 mA with Ni-filtered Cu Kα radiation (λ = 1.54056 Å), and scanning electron microscopy (SEM) (QUANTA FEG 450, FEI Company, Eindhoven, The Netherlands) were used to determine the surface morphologies. The as prepared silver sols were centrifuged at 10 000 rpm for 20 minutes at room temperature and the solid AgNPs were obtained and then blended with KBr powder and pressed into a pellet for FTIR measurements.

2.3 Encapsulation studies

The reversible encapsulation of starch-capped AgNPs was analyzed by iodimetric titrations. In a separate experiment, AgNPs were prepared by adding AgNO₃ (5.0 cm³; 0.01 mol dm⁻³) into a solution containing starch (2.0 cm³; 2.0% w/v) and [extract] = 33.3% v/v. The resulting brownish silver sol (10 cm³) was titrated against 0.1 N iodine solution (I₂/KI reagent) and a deep blue colour of the starch–iodine complex was obtained. 3.0 cm³ of AgNP solution was further added into the blue colour complex solution and UV-visible spectra were recorded at regular intervals. The morphology of the blue colour complex and the sol obtained after the reversible encapsulation of AgNPs was also confirmed by TEM measurements.

3 Results and discussion

3.1 Extract coated AgNPs

Water-soluble small metal particles have been found to be advantageous over the water-insoluble forms because UV-visible spectrophotometric methods can be used to monitor the optical changes that accompany the surface reactions (solutions of nanometer large particles are transparent and the scattering of light can be neglected).³² It is well known that surface plasmon resonance bands are influenced by the size, shape, composition, and dielectric environment of the reaction medium.³³ The shape of the UV-visible spectra and position of the maximal wavelength gives initial information about the size and the size distribution of the Ag-nanoparticles.³⁴ The most characteristic part for AgNPs is a narrow plasmon absorption band observable in the 350–600 nm region. In order to establish the role of D. deltoidea extract as a reducing and stabilizing agent in the synthesis of AgNPs, a series of experiments were performed as a function of [Ag⁺] (=3.3 × 10⁻⁴ to 23.3 × 10⁻⁴ mol dm⁻³) at constant [extract] (=33.3% v/v) and temperature (=30 °C). The effects of [extract] (from 3.3 to 23.3% v/v) were also studied at fixed [Ag⁺] (=16.6 × 10⁻⁴ mol dm⁻³) and temperature (=30 °C). These observed results are given in Fig. S1A and B (ESI†) for the variation of [Ag⁺] and [extract], respectively. The optical images of the Ag⁺ ions only and Ag-nanoparticles are shown in Fig. S2 (ESI†) as a function of time, indicating that the typical color of the Ag-nanoparticles changed from pale yellow to yellow, orange, light red, and wine red with time. The optical images of the Dioscorea deltoidea tubers, their aqueous extract, and the formation of AgNPs after the addition of AgNO₃ into the extract are summarized in Scheme 1 as a flow diagram showing the systematic step-wise presentation to the formation of AgNPs.

Scheme 1 Optical images of Dioscorea deltoidea tubers, their aqueous extract, and AgNPs.

The morphology (size, shape and size distribution) and crystalline nature of the Ag-nanoparticles (reaction conditions: [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, [extract] = 33.3% v/v, temperature = 30 °C) were determined by using TEM images and selected area electron diffraction ring pattern measurements (Fig. 1). Interestingly, mainly truncated triangular nano plates along with some nano-rod, spherical, hexahedral, and highly poly-dispersed particles are observed. The average edge length and the thickness of the truncated triangular polyhedral nanoplates were ca. 11.2 nm and 98.9 nm, respectively. The nano-disks could be formed by the dissolution of the corner atoms of the truncated triangular nano-plates (Fig. 1(A)). The single-crystalline nature of the truncated triangular particles was also confirmed by selected area electron diffraction ring patterns which indicated that the nanoplates were oriented with {111} planes as the basal plane (Fig. 1(B)). A thin layer of another material was also seen on the surface of the Ag-nanoparticles, which might be due to the capping properties of the organic materials of the extract. The main constituent, diosgenin, of D. deltoidea tuber extract was responsible for the reduction of Ag⁺ ions into metallic Ag⁰. Hydroxy groups of steroidal diosgenin reduce Ag⁺ into Ag⁰ (rate-determining step, rds). The metallic Ag⁰ reacts with Ag⁺ to form Ag₂⁺, which undergoes dimerization to yield Ag₄²⁺ (yellow-color silver sol; stable species for a long time even under air and growth stops at the stage of this species). The capping and/or shape-directing role may be explained in terms of excess diosgenin adsorption onto the surface of the Ag-nanoparticles. The above explanations and proposed mechanism are in good agreement with the hypothesis developed by Mahal et al.³⁵ and Wu et al.³⁶ regarding the reducing-cum-stabilizing role of proteins in the synthesis of silver and gold nanoparticles.

Fig. 1 TEM images (A) and selected electron diffraction ring patterns (B) of AgNPs. Reaction conditions: [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, and [extract] = 33.3% v/v.

3.2 Starch coated Ag-nanoparticles

The use of environmentally benign and renewable materials like water soluble starch offers numerous benefits in terms of eco-friendliness and compatibility with pharmaceutical and biomedical applications in nanotechnology. In order to prepare the starch capped Ag-nanoparticles and to find a suitable stabilizer, the effect of a biodegradable natural polymer, [starch], was also studied on the appearance of characteristic yellowish-red color silver sols. An intense yellow color appeared within a short period of time in the presence of water soluble starch indicating that the typical color of the Ag-nanoparticles changed from pale yellow to yellow, orange, light red, and wine red with time. The appearance and/or change of color strongly depend on the presence of starch in the reaction mixture (Fig. 2). The morphology of the Ag-nanoparticles can be controlled not only by the starch but also by reaction-time alteration. From these observations it is clear that starch not only stabilized and/or capped the nanoparticles but also altered the nucleation process (Fig. S2† and 2). We do not see any marked difference in the overall nature of the spectrum, shape and size of the extract- and starch-capped AgNPs produced. On the other hand, we observed a significant difference in the optical images of the AgNPs prepared with the aqueous extract of Dioscorea deltoidea in the absence and presence of starch (Fig. 2 and S2†), which might be due to the fast adsorption of AgNPs into the helix chain of amylose through electrostatic interactions between the positive surface of the AgNPs and the lone-pair electrons of the –OH groups. Anisotropy among the AgNPs could be due to the reasons given by Bakshi et al. that metallic nanocrystals formed using zein protein were gradually enclosed by protective molecules, which eliminated rapid sintering of smaller nanoparticles leading to the formation of nanoparticles having mixed morphologies.³⁵

Fig. 2 Optical images of Ag⁺ ions only (A) and AgNP formation in the presence of starch (2.0 cm³; 2.0% w/v) at different time intervals. Reaction conditions: [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, [extract] = 33.3% v/v, and time = 10 (B), 30 (C), 90 (D) and 180 min (E).

To determine the optimum [starch] required for the stabilization of the Ag-nanoparticles, a series of experiments were carried out under different [starch] (range: from 2.0 cm³; 2.0% w/v to 6.0 cm³; 2.0% w/v) at fixed [Ag⁺] (16.6 × 10⁻⁴ mol dm⁻³), and [extract] (33.3% v/v). The absorbance remained constant with increasing [starch] and was found to be 0.31, 0.30, 0.30 and 0.31 for 2.0, 3.0, 4.0, and 5.0 cm³ [starch], respectively. Addition of a small quantity of starch has a pronounced effect on the reaction path of Ag-nanoparticle formation. A small concentration of starch ≥2.0 cm³; 2.0% w/v is enough to catalyze and/or stabilize the nanoparticles. Fig. S3A and B show (ESI†) the UV-visible spectra of the Ag-nanoparticles prepared at constant [extract] = 33.3% v/v, starch = 2.0 cm³; 2% w/v, temperature = 30 °C and [Ag⁺] varied from 3.3 to 33.3 × 10⁻⁴ mol dm⁻³. The formation of Ag-nanoparticles was confirmed by the appearance of the characteristic SPR band at 425 nm. On increasing the [Ag⁺], the absorbance increased and a sharp peak developed at higher [Ag⁺]. Fig. 1SB† shows the effects of [extract] on the UV-visible spectra of Ag-nanoparticles obtained at constant [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, starch (2.0 cm³; 2% w/v), temperature = 30 °C, and [extract] varied from 3.3 to 33.3% v/v. The peak wavelength did not shift significantly during the reaction with increasing [extract] indicating that the morphology of the Ag-nanoparticles did not strongly depend on the [extract]. As the reaction proceeded the width of the band also increased which might be due to the excitation of different multiple modes present in the faceted and anisotropic growth of the particles. At higher [Ag⁺] ≥ 33.3 × 10⁻³ mol dm⁻³, the reaction mixture became turbid instead of a perfectly transparent silver sol which might be due to the unlimited and/or uncontrolled growth of the AgNPs. The appearance of a broad band with [starch] indicates that the initially reduced AgNPs grow to form larger particles and finally starch acts as a shape-directing agent. At higher [extract] more than 33.3% v/v, aggregation occurs and the formed AgNPs precipitated. So the most suitable concentration for AgNP preparation is ≤23.3% v/v [extract], which is suitable for industrial applications.

To check the stability of the resulting silver sols, the absorbance at 425 nm was monitored at an interval of 24 h. No significant change in absorbance was noticed indicating the formation of starch-capped Ag-nanoparticles. To ascertain the presence of Ag⁺ ions, aqueous solutions of NaCl, NaBr and NaI (5.0 cm³; 0.01 mol dm⁻³) were added to the starch-capped AgNP solution after completion of the reaction. Interestingly, the formation of AgCl, AgBr and AgI precipitates were not observed in the presence of NaCl, NaBr and NaI, indicating that there were no free Ag⁺ ions in the starch-capped AgNP solution. On the other hand, we observed the appearance of white and yellowish turbidity after the addition of NaCl, NaBr and NaI in an aqueous AgNO₃ (5.0 cm³; 0.01 mol dm⁻³) solution. Reaction–time plots suggest that there was no significant synthesis after a short reaction time of ca. 30 min. The reaction-rate increased with time very rapidly for 2 h and the reaction was completed within 3 h (Fig. S4†). Evidently new particles were not formed: the process of nucleation may be regarded as finished since the absorbance remained nearly constant.

Fig. 3 shows the TEM images and selected area electron diffraction ring patterns of the starch-capped Ag-nanoparticles. From these results it is evident that the particles are aggregated, capped and/or encapsulated by starch. The reason for the aggregation might be due to the presence of the helix like structure of amylose (main constituent of water soluble starch). The TEM image of Fig. 3(A) also reveals that these Ag-nanoparticles are aggregated and/or deposited inside the water soluble helical chain of amylose. We did not observe this type of aggregation without starch (Fig. 1(A)). Diffraction rings can be seen when the corresponding selected area electron diffraction of the Ag-nanoparticles was conducted (Fig. 3(B)). The six-fold symmetry of the diffraction spots indicates that the surface of the particles was bounded by {111} faces. The other sets of spots could be identified as the {220}, {311}, {420} and {440} planes indicating the formation of a pure face-centered cubic (fcc) silver structure. These results are in good agreement with the observations of Bakshi and his co-workers (2011 and 2013) that the capping ability of lipids and/or normal surfactants originates from the electrostatic interactions between the polar head groups and charged nanoparticle surface. Surface passivation exists in the form of a lipid bilayer which provides excellent steric and charge stabilization (due to Derjaguin–Landau–Verwey–Overbeek theory) to the colloidal nanoparticles. Nano-spheres dominated the population of the synthesized silver nanoparticles as shown in the TEM images and in some cases anisotropy was also observed similar to other reports. These results are in good agreement with the observations of Raveendran et al. regarding the capping behaviours of polysaccharide functionalized AgNPs through –OH groups acting as a soft template.^14,15

Fig. 3 TEM image (A) and selected electron diffraction ring pattern (B) of AgNPs in the presence of starch (2.0 cm³; 2%). Reaction conditions: [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, and [extract] = 33.3% v/v.

Sun and Xia reported a shape-controlled synthesis of silver nanocubes in large quantities by reducing silver nitrate with ethylene glycol in the presence of poly(vinylpyrrolidone) and used the resulting nanocubes as sacrificial templates to form gold nanoboxes with a well-defined shape and hollow structure. The morphology of the as prepared silver nanocubes were found to strongly depend on reaction conditions (temperature, [AgNO₃], and the molar ratio between the repeating unit of capping agent and AgNO₃). They suggested that the major requirement for the shape-controlled synthesis of metal nanoparticles seemed to be the selection of an appropriate capping reagent.³⁷ Xiao and Qi in their feature article summarized the surfactant assisted shape controlled synthesis of gold nanocrystals and pointed out that the morphology of branched gold nano-crystals (stars, flowers, combs, and dendrites) depends on the method and nature of the capping agents.³⁸ We obtained mostly triangular truncated AgNPs of 11.2 to 87.6 nm with no drastic change in the shape and size in the absence (Fig. 1) and presence of starch (Fig. 3(A)). The hydrophobic nature of steroidal diosgenin and lack of appropriate surface adsorption allows the growing nuclei to grow into independent AgNPs without any kind of self-aggregation and the growth is much slower in comparison with that in the presence of starch (Fig. 2 and S2:† optical images). Thus we may state confidently that the control of morphology is a practical reality, which cannot be controlled by a single parameter.^37–40

To determine the surface properties and composition of the AgNPs, the resulting products were collected by centrifugation, washed with double distilled water, and re-dispersed in water for use. SEM, XRD, and EDX of the nanoparticles are depicted graphically in Fig. 4 and 5, respectively. Fig. 4(A) shows the SEM image of the high-density, mostly triangular nanoplates and highly poly-dispersed and relatively spherical AgNPs. The SEM image of the silver nanoparticles shows electrostatic interactions between the capping molecules bound to the AgNPs. Inspection of these results suggests that the nanoparticles are adsorbed on the surface of the amylose helix and the various layered-like structures of starch. The XRD peak positions (Fig. 4(B)) are consistent with metallic silver. Bragg’s reflections are observed in the XRD pattern at 2θ values of 32.2, 38.1, 44.6, 64.04, and 77.30, which correspond to the Miller indices (111), (200), (220), (311) and (222), respectively. Energy dispersive X-ray analysis used particularly for the determination of elemental composition, was also performed to estimate the elemental composition and purity of the AgNPs. EDX analysis of the particles revealed a strong signal for silver at 3.2 keV, characteristic of pure silver nanoparticles (Fig. 5), which is confirmation of the composition of >99% of Ag metal atoms in the starch-coated nanoparticles.

Fig. 4 SEM image (A) and XRD pattern (B) of starch-capped AgNPs. Reaction conditions: [extract] = 33.3% v/v, [AgNO₃] = 10 × 10⁻⁴ mol dm⁻³ and starch = 2 cm³ of 2% starch solution.

Fig. 5 Energy-dispersive X-ray spectroscopy of the AgNPs. Reaction conditions: [extract] = 33.3% v/v, [starch] = 2 cm⁻³; 2.0% w/v and [AgNO₃] = 16.6 × 10⁻⁴ mol dm⁻³.

FT-IR spectroscopy is a useful tool to determine and/or confirm the functional groups of any organic molecule. Fourier-transform infrared (FT-IR) analysis was carried out on an IRPrestige-21, IRAffinity-1, FTIR-8400S (Shimadzu Corporation Analytical and Measuring Instrument Division) using a ZnSe cell at room temperature, in the wavelength range of 4000–500 cm⁻¹. The FT-IR spectra of Dioscorea deltoidea tuber extract before and after reduction of silver are shown in Fig. 6(A and B). The absorption band at 3325 cm⁻¹ is due to OH groups present in steroidal sapogenin. The peaks obtained at 1650 cm⁻¹ and 1410 cm⁻¹ are due to associated water and the stretch vibrations of alkyl groups Fig. 6(A). A little blue shift was observed in the peak positions of Fig. 6(B) as compared to Fig. 6(A). The intensity of the broad band around 3334 cm⁻¹ in the extract, Fig. 6(A), started to decrease upon addition of the extract into silver sol as shown in Fig. 6(B). This indicates that steroidal sapogenin is mainly responsible for reduction of Ag⁺ into AgNPs. Further, the two new peaks appearing after addition of the extract into the silver sol at 2945 cm, 2850 cm, in Fig. 6(B), could be assigned to the –CH stretching vibrations of the –CH₃ and –CH₂ functional groups. In Fig. 6(C) peaks at 3440 cm⁻¹, 2945 cm⁻¹, 1036 cm⁻¹, and 900–1200 cm⁻¹ are due to O–H stretching, C–H stretching, C–OH stretching, and glycosidic linkage, respectively, and are attributed to free starch. While in Fig. 6(D) peaks around 3341 cm⁻¹, 2916 cm⁻¹, 2865 cm⁻¹, and 1655 cm⁻¹ correspond to hydroxyl groups, C–H stretching absorption bands of alkane and the stretching vibration of alkyl groups.

Fig. 6 FT-IR spectra of extract only (A), extract-capped AgNPs (B), pure starch (C) and starch-capped AgNPs (D). Reaction conditions: [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, [extract] = 33.3% v/v and [starch] = 2.0 cm³ of 2.0% w/v (D), temperature = 26 °C.

3.3 Encapsulation studies

It has been established that micelles, vesicles, and helical structures of starch with controllable sizes help the encapsulation of guest molecules with intended applications in solubilization, catalysis, controlled delivery and growing materials with the structural motif of the host due to the presence of hydrophobic molecules inside their hydrophobic core while the outer hydrophilic layer helps in dissolving in water.⁴¹ Amylose (water soluble constituent of starch) is known to form helical structures of variable pocket sizes according to the size of the host. The structure of amylose–iodine deep-blue complex is well known.^42,43 Yu et al. re-examined the structure of this complex by using different techniques such as Raman spectroscopy, UV-visible and second-derivative UV-visible spectroscopies.⁴⁴ They reported that the shape of the UV-visible spectra and position of the absorption maxima (λ_max = 480–510, 610–640, 690–720, and 730–760 for I₉⁻, I₁₁⁻, I₁₃⁻, and I₁₅⁻, respectively) depend on the subunits of the polyiodide chains (I₃⁻ and I₅⁻) which exist inside the helix of amylose. Stoddart et al. used amylose–iodine blue complex to solubilize as well as purify single-walled carbon nanotubes (insoluble in an aqueous solution of starch) in water. They recorded the UV-visible spectra of amylose–iodine complex and the amylose–carbon nanotube complex obtained from the blue amylose–iodine complex in water and suggested that the blue color was lost upon solubilization of the carbon nanotubes in an aqueous solution of amylose.⁴⁵ Sarma and Chattopadhyay reported a method for the complete reversible encapsulation of polyaniline and a polyaniline–gold-nanoparticle composite in starch by replacement with molecular iodine.¹⁶ Therefore, to obtain insight into the encapsulation of starch capped Ag-nanoparticles, 0.10 N KI–I₂ reagent was prepared.

For this, the required amount of iodine was dissolved in water containing the requisite potassium iodide solution because iodine is not very soluble in water. This makes an iodide–starch complex and leads to the formation of an intense blue-black color. Teitelbaum et al. used Resonance Raman and Mossbauer techniques for elucidating the structure of the deep-blue amylose–iodine complex and suggested that the pentaiodide moiety of the KI–I₂ reagent was encapsulated within the amylose helix.⁴⁶ Starch amylopectin does not produce any complex with KI–I₂. The UV-visible absorption spectrum of the yellowish-brown colored silver sols (starch capped Ag-nanoparticles), KI–I₂ reagent (alone), starch-encapsulated Ag-nanoparticles and also that of the starch–I₂ blue complex were recorded (Fig. 7). The absorption spectrum of the starch-capped Ag-nanoparticles consists of a sharp band at 425 nm (Fig. 7 (■)) while the KI–I₂ regent (light brown color) covers the whole visible region of the spectrum (Fig. 7(♦)). Upon addition of KI–I₂ (0.10 cm³; 0.10 N) to a solution of starch-capped yellow colored silver sol (10.0 cm³; [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³), the solution becomes pale yellow. The peak at 425 nm disappears completely and a new shoulder begins to develop at ca. 400 nm (Fig. 7 (○)) which might be due to the formation of poly-dispersed Ag-nanoparticles.

Fig. 7 Spectra of KI–iodine reagent (♦), and starch capped Ag-nanoparticles (■). Starch capped nanoparticle solution was titrated with KI–iodine, and the UV-vis spectra at various stages are as follows: (yellow turbidity; ○) initial stage indicating the morphological changes, formation of the amylose–iodine complex (●) and the addition of excess silver nanoparticles resulting in the reversible encapsulation of Ag-nanoparticles (▼). Optical images of AgNPs (A), blue complex (B) and reversible encapsulated nanoparticles (C). Reaction conditions: [Ag⁺] = 16.6 × 10⁻⁴ mol dm⁻³, [extract] = 33.3% v/v, [starch] = 2.0 cm³ of 2.0% w/v, temperature = 30 °C.

Upon further addition of KI–I₂ reagent (0.05 cm³; 0.1 N), the same solution (pale yellow color) becomes deep-blue and shows two peaks at 400 nm and 625 nm (Fig. 7 (●)), which might be due to the encapsulation of polyiodide subunit chains of I₁₁⁻ into the helix of amylose.⁴⁴ The stoichiometry of the complexation between KI–I₂ and starch (deep blue color) present on the surface of the AgNPs was calculated and found to be [starch] = 0.13% w/v and KI–I₂ = 14.7 × 10⁻⁴ mol dm⁻³ for [AgNPs] = 15.7 × 10⁻⁴ mol dm⁻³. The blue color as well as the UV-visible peak due to starch–I₂ vanished in an excess of AgNPs (3.0 cm³ of 16.6 × 10⁻⁴ mol dm⁻³). The small [AgNPs] (=3.3 × 10⁻⁴ mol dm⁻³) was responsible for diminishing the peak of the blue complex. Finally, the reaction mixture shows only one peak at 400 nm (Fig. 7 (▼)). It thus seems that the AgNPs are rather unstable in the presence of KI–I₂ reagent and are rapidly converted into larger particles, which might be due to the reversible encapsulation of the Ag-nanoparticles. On the basis of the above observations, we conclude that starch capped Ag-nanoparticles are encapsulated in much the same way as I₂ is known to be encapsulated in starch. The red-shift is attributed to the strong interactions between the adsorbed Ag⁺ and iodide ions on the surface of Ag₄²⁺. The optical images also showed that the typical color of the starch capped Ag-nanoparticles changed from yellowish orange (starch capped Ag-nanoparticles; Fig. 7(A)), to deep blue (starch–KI–iodine complex; Fig. 7(B)) and pale yellow (reversible encapsulation of silver nanoparticles; Fig. 7(C)). Our observations are in good agreement with the reversible encapsulation of polyaniline, composites, and single walled carbon nano tubes, which could be recovered in water by replacement with molecular iodine.^16,44

In order to confirm the results of Fig. 7, TEM images were also recorded for each titration. TEM analysis shows that the morphology of the particles changed drastically in the presence of KI–I₂ reagent. The size of the blue starch–iodine complex nanoparticles is very small as compared to the size of the starch capped AgNPs and also the shape changes from triangular to rod and hexagonal irregular nanoparticles (Scheme 1). After the addition of excess AgNPs into the resulting deep-blue colored complex, yellowish turbidity appeared and smaller particles aggregated leading to the formation of beautiful branched-like AgNPs (TEM images; Scheme 2). It thus seems that the small AgNPs are rapidly converted into larger particles, which might be due to the reversible encapsulation.

Scheme 2 TEM images of reversible encapsulation starch capped Ag-nanoparticles.

On the basis of Scheme 2, we may state confidently that AgNPs were being encapsulated in much the same way as molecular iodine is known to be encapsulated in starch.⁴⁴

To see whether the aqueous extract was capable of the encapsulation of AgNPs under our experimental conditions; some other experiments were also performed. Interestingly, we did not observe the formation of the deep blue color complex with KI–I₂ in a constant volume (10.0 cm³) of extract capped Ag-nanoparticles. This is not surprising since blue complex formation is a characteristic reaction of amylose and KI–I₂ reagent. This suggests that the constituent(s) of Dioscorea deltoidea acted as a reducing, stabilizing and/or capping agent. They did not behave like a supramolecular assembly for the encapsulation of guest molecules. On the basis of the above results, we propose that the AgNPs were being encapsulated in much the same way as molecular iodine is known to be encapsulated in starch (Scheme 3).

Scheme 3 Schematic representations of the different behaviours of extract-, and starch-capped AgNPs.

Nanoencapsulation depends on the choice of a suitable polymeric system having maximum encapsulation (higher encapsulation efficiency). Chu and his co-workers reported a method for the synthesis of glucose-sensitive polyamide microcapsules having a porous membrane and functional gates.⁴⁷ Wang et al. in their pioneering review, pointed out that microparticles with a hydrogel or porous shell offer excellent performance for encapsulation and controlled release.⁴⁸ It has been established that the functional gates in membrane pores (linear polymer chains, cross linked hydrogel networks, and microspheres) were responsible for drug delivery.^49–51 Dioscorea deltoidea has also been used as a raw material for the synthesis of steroidal drugs. Thus we may state confidently that the encapsulation and/or release of AgNPs might be due to the helical shape of amylose.⁵¹ Our results (starch-capped encapsulated AgNPs) could deliver nanoparticles in the human body to cure the infections and diseases. Amylose (water soluble) can easily be detached from the surface of AgNPs which could be used as a drug release vehicle for biomedical and other pharmaceutical applications in the future.

4 Conclusions

In this study we reported the reduction of silver nitrate by diosgenin, a steroid sapogenin in the absence and presence of water soluble starch. The addition of NaCl and NaBr in the silver sol does not lead to precipitate formation indicating the reduction of all Ag⁺ into Ag⁰. TEM images indicated the formation of spherical, triangular, rod shaped, hexagonal, poly-dispersed and aggregated Ag-nanoparticles in the presence and absence of starch, respectively. An interesting observation was the formation of a deep blue complex, and a decrease in size and distribution upon the addition of KI–iodine reagent into the starch capped Ag-nanoparticles. It was also reported that encapsulated silver nano-composites in starch could be recovered by replacement with molecular iodine. Optical images reveal the overall role and aggregation behavior of AgNPs and demonstrate how effectively starch controls the nucleation and aggregation through fusogenic behavior. Due to the presence of a large number of –OH groups in amylose, it readily adsorbs on the positive AgNP surface and thus generates bio-conjugated nanoparticles suitable for drug delivery in the human body.

References

1. A. Henglein, Chem. Rev., 1989, 89, 1861–1873.
2. M. Poliakoff and P. Anastas, Nature, 2001, 413, 257–258.
3. J. L. Gardea-Torresdey, E. Gomez, J. R. Peralta-Videa, J. G. Parsons, H. Troiani and M. Jose-Yacaman, Langmuir, 2003, 19, 357–1361.
4. M. Grzelczak, J. Perez-Juste, P. Mulvaney and L. M. Liz-Marzan, Chem. Soc. Rev., 2008, 37, 1783–1791.
5. M. S. Bakshi, J. Phys. Chem. C, 2011, 115, 13947–13960.
6. H. Huang and X. Yang, Carbohydr. Res., 2004, 339, 2627–2631.
7. B. Nikoobakht and M. A. El-Sayed, Langmuir, 2001, 17, 6368–6374.
8. H. Xie, J. Y. Lee, D. I. C. Wang and Y. P. Ting, ACS Nano, 2007, 1, 429–439.
9. P. Khullar, V. Singh, A. Mahal, P. N. Dave, S. Thakur, G. Kaur, J. Singh, S. S. Kamboj and M. S. Bakshi, J. Phys. Chem. C, 2012, 116, 8834–8843.
10. Z. Khan and A. Y. Obaid, RSC Adv., 2016, 6, 29116–29126.
11. M. Amanullah and L. Yu, J. Pet. Sci. Eng., 2005, 48, 199–208.
12. V. K. Sharma, R. A. Yngard and Y. Lin, Adv. Colloid Interface Sci., 2009, 145, 83–96.
13. Z. Shervani and Y. Yamamoto, Carbohydr. Res., 2011, 346, 651–658.
14. P. Raveendran, J. Fu and S. L. Wallen, J. Am. Chem. Soc., 2003, 125, 13940–13941.
15. P. Raveendran, J. Fu and S. L. Wallen, Green Chem., 2006, 8, 34–38.
16. T. K. Sarma and A. Chattopadhyay, Langmuir, 2004, 20, 3520–3524.
17. K. Esumi, T. Hosoyo, A. Yamahira and K. Torigoe, J. Colloid Interface Sci., 2000, 226, 346–352.
18. Y. Badr and M. A. Mahmoud, J. Appl. Polym. Sci., 2006, 99, 3608–3614.
19. M. H. El-Rafie, M. E. El-Naggar, M. A. Ramadan, M. M. G. Fouda, S. S. Al-Deyabc and A. Hebeish, Carbohydr. Polym., 2011, 86, 630–635.
20. P. Dallas, V. K. Sharm and R. Zboril, Adv. Colloid Interface Sci., 2011, 166, 119–135.
21. N. Vigneshwaran, R. P. Nachane, R. H. Balasubramanya and P. V. Varadarajan, Carbohydr. Res., 2006, 341, 2012–2018.
22. Z. Khan, T. Singh, J. I. Hussain, A. Y. Obaid, S. A. Al-Thabaiti and E. H. El-Mossalamy, Colloids Surf., B, 2013, 102, 578–584.
23. O. Bashir, S. Hussain, Z. Khan and S. A. Al-Thabaiti, Carbohydr. Polym., 2014, 107, 167–173.
24. K. Vimala, K. S. Sivudu, Y. M. Mohan, B. Sreedhar and K. Raju, Carbohydr. Polym., 2009, 75, 463–471.
25. D. G. Yu, Colloids Surf., B, 2007, 59, 171–178.
26. J. W. Kim, J. E. Lee, J. H. Ryu, J. S. Lee, S. J. Kim and S. H. Han, et al., J. Polym. Sci., Part A: Polym. Chem., 2004, 42, 2551–2554.
27. J. W. Rhim, S. I. Hong, M. Park and P. K. W. Ng, J. Agric. Food Chem., 2006, 54, 5814–5822.
28. A. Kumari, S. K. Yadav and S. C. Yadav, Colloids Surf., B, 2010, 75, 1–18.
29. Z. Zaheer and Rafiuddin, Colloids Surf., B, 2013, 108, 90–94.
30. E. S. Aazam and Z. Zaheer, Bioprocess Biosyst. Eng., 2016, 39, 575–584.
31. S. S. Shankar, A. Ahmad and M. Sastry, Biotechnol. Prog., 2003, 19, 1627–1631.
32. A. Henglein, P. Mulvaney and T. Linnert, Faraday Discuss., 1991, 92, 31–44.
33. M. S. Bakshi, J. Phys. Chem. C, 2011, 115, 13947–13960.
34. D. W. Kim, S. I. Shin, and S. G. Oh, in Surfactant Sciences Series, Marcel Dekker, ed. K. L. Mittal and D. O. Shah, New York Basel, 2003.
35. A. Mahal, P. Khullar, H. Kumar, G. Kaur, N. Singh, M. Jelokhani-Niaraki and M. S. Bakshi, ACS Sustainable Chem. Eng., 2013, 1, 627–639.
36. H. Wu, L. He, M. Gao, S. Gao, X. Liao and B. Shi, New J. Chem., 2011, 35, 2902–2909.
37. Y. Sun and Y. Xia, Science, 2002, 298, 2176–2179.
38. J. Xiao and L. Qi, Nanoscale, 2011, 3, 1383–1396.
39. T. S. Ahmadi, Z. L. Wang, T. C. Green, A. Henglein and M. A. El-Sayed, Science, 1996, 272, 1924–1929.
40. R. Jin, Y. W. Cao, C. A. Mirkin, K. L. Kelly and G. C. Schatz, Science, 2001, 294, 1901–1903.
41. D. Balasubramanian, B. Raman and C. V. Sundari, J. Am. Chem. Soc., 1993, 115, 74–77.
42. R. E. Rundle, J. Am. Chem. Soc., 1947, 69, 1769–1772.
43. C. A. Knutson, Carbohydr. Polym., 1999, 42, 65–72.
44. X. Yu, C. Houtman and R. H. Atalla, Carbohydr. Res., 1996, 292, 129–141.
45. A. Star, D. W. Steuerman, J. R. Heath and J. F. Stoddart, Angew. Chem., Int. Ed., 2002, 41, 2508–2512.
46. R. C. Teitelbaum, S. L. Ruby and T. J. Marks, J. Am. Chem. Soc., 1978, 100, 215–3217.
47. L.-Y. Chu, Y.-J. Liang, W.-M. Chen, X.-J. Ju and H.-D. Wang, Colloids Surf., B, 2004, 37, 9–14.
48. W. Wang, M.-J. Zhang and L.-Y. Chu, Acc. Chem. Res., 2014, 47, 373–384.
49. J. Wei, X.-J. Ju, X.-Y. Zou, R. Xie, W. Wang, Y.-M. Liu and L.-Y. Chu, Adv. Funct. Mater., 2014, 24, 3312–3323.
50. D. Menne, F. Pitsch, J. E. Wong, A. Pich and M. Wessling, Angew. Chem., Int. Ed., 2014, 53, 5706–5710.
51. Z. Liu, W. Wang, R. Xie, X.-J. Ju and L.-Y. Chu, Chem. Soc. Rev., 2016, 45, 460–475.

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Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra09319a

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