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  • Research
  • Open Access

Biofibres from biofuel industrial byproduct—Pongamia pinnata seed hull

  • 1Email author,
  • 1 and
  • 1
Bioresources and Bioprocessing20174:14

https://doi.org/10.1186/s40643-017-0144-x

©  The Author(s) 2017

  • Received: 15 December 2016
  • Accepted: 8 February 2017
  • Published:

Abstract

Background

Biodiesel production using Pongamia pinnata (P. pinnata) seeds results in large amount of unused seed hull. These seed hulls serve as a potential source for cellulose fibres which can be exploited as reinforcement in composites.

Methods

These seed hulls were processed using chlorination and alkaline extraction process in order to isolate cellulose fibres. Scanning electron microscopy (SEM), dynamic light scattering (DLS), thermogravimetric analysis (TGA), X-ray diffraction (XRD), Fourier transform infrared spectroscopy (FTIR) and nuclear magnetic resonance spectroscopy (NMR) analysis demonstrated the morphological changes in the fibre structure.

Results

Cellulose microfibres of diameter 6–8 µm, hydrodynamic diameter of 58.4 nm and length of 535 nm were isolated. Thermal stability was enhanced by 70 °C and crystallinity index (CI) by 19.8% ensuring isolation of crystalline cellulose fibres.

Conclusion

The sequential chlorination and alkaline treatment stemmed to the isolation of cellulose fibres from P. pinnata seed hull. The isolated cellulose fibres possessed enhanced morphological, thermal, and crystalline properties in comparison with P. pinnata seed hull. These cellulose microfibres may potentially find application as biofillers in biodegradable composites by augmenting their properties.

Keywords

  • Cellulose microfibres
  • Pongamia pinnata seed hull
  • Hemicellulose
  • Lignin
  • Chlorination

Background

Cellulose is nature’s most lavishly available polymer. Highly purified cellulose fibre is been isolated from several plant sources, such as branch barks of mulberry (Li et al. 2009), pineapple leaf fibres (Cherian et al. 2010; Mangal et al. 2003), pea hull fibre (Chen et al. 2009), coconut husk fibres (Rosa et al. 2010), banana rachis (Zuluaga et al. 2009), sugar beet (Dinand et al. 1999; Dufresne et al. 1997), wheat straw (Kaushik and Singh 2011), palm leaf sheath (Maheswari et al. 2012), Arundo donax L stem (Fiore et al. 2014), cotton stalk (Hou et al. 2014).

From the past two decades these biofibres are being used as filler material in the preparation of composites and have gained prodigious attention (Hubbe et al. 2008). In view of better utilization of renewable resources, there is a need to explore other renewable greener sources, which can be utilized in developing high strength light weight biocomposites for high-end applications. Pongamia pinnata seed hull is chosen for the present work to exploit its potential for cellulose fibres which could be utilized as reinforcement in biocomposites. In India and south East Asia, Pongamia pinnata (Karanja) seed is used for biodiesel production (Demirbas 2009). It is also a traditional medicinal plant with all parts having certain medicinal value (Yadav et al. 2004). Biofuel production using P. pinnata seeds has resulted in large-scale cultivation of these trees (Shwetha et al. 2014). The biofuel processing fallouts in significant amount of residual P. pinnata seed hull, in which cellulose percentage approximates to 40% and is similar as in shelly wood (Nadeem et al. 2009). Thus these underused seed hulls can find potential application as a source for cellulose fibres.

Isolation of cellulose fibres is customarily carried out by mechanical treatments such as homogenisation (Du et al. 2016; Julie et al. 2016), sonication, (Sheltami et al. 2012; Saurabh et al. 2016), steam explosion (Saelee et al. 2014) etc.; chemical treatments such as acid hydrolysis (Abidin et al. 2015), TEMPO oxidation (Du et al. 2016), chlorination and alkaline treatments (Sheltami et al. 2012; Johar et al. 2012; Maheswari et al. 2012) etc.; enzymatic treatments (Saelee et al. 2014) and conjointly with the combination of two or more of the aforementioned processes. Chemical treatments usually act upon the binding material of the fibril structure enabling the fibres to individualize (Johar et al. 2012). Chlorination treatment being a chemical treatment is a well-established treatment which assists isolation of high quality pure cellulose fibres by bleaching and delignifies the cellulose material, while alkali treatment dissolves the wax, pectin and hemicellulose ensuring efficient isolation of cellulose microfibres. These chemical methods are used in combination to isolate cellulose fibres from different sources (Espino et al. 2014; Johar et al. 2012; Sheltami et al. 2012; Mandal and Chakrabarty 2011; Moran et al. 2008) and are also found to be efficient and economical when compared to high energy-consuming mechanical methods (Motaung and Mtibe 2015).

In the present research work, the cellulose fibres were isolated from the P. pinnata seed hull using chlorination and alkaline process. The isolated cellulose microfibres were characterized using scanning electron microscopy (SEM), dynamic light scattering (DLS), thermogravimetric analysis (TGA), X-ray diffraction (XRD), Fourier transform infrared spectroscopy (FTIR) and nuclear magnetic resonance spectroscopy (NMR) analysis for their morphological, thermal and crystalline properties.

Methods

Materials

Pongamia pinnata seed hulls were collected from “SEEDS” Research Centre, University of Agricultural Sciences, Bengaluru, India. All the chemicals used were of analytical grade.

Fibre processing

Pongamia pinnata seed hulls were separated from stones and other plant materials by hand picking. The dust and mud particles sticking to the seed hulls were removed by washing them extensively in tap water and finally with distilled water. Later dried under sunlight for two days and stored in sealed polythene bags for further use. Cleaned seed hulls were ground, screened (0.25 mm sieves) and oven dried at 105 °C for 8 h.

Isolation of cellulose microfibres

Cellulose microfibres were isolated from P. pinnata seed hull by chlorination and alkaline extraction process (Maheswari et al. 2012). Cleaned seed hull fibres were dewaxed using toluene–ethanol mixture (2:1) for 6 h. Excess of solvent from the fibres was removed by suction and later kept for drying in hot air oven. Fibres were bleached with 7% NaClO2 taken in fibre to liquor ratio of 1:50 (pH vicinity 4–4.2 was maintained using acetic acid and sodium acetate buffer) for 2 h at 100 °C and was washed successively using 2% sodium bisulphate, distilled water and ethanol. Further the extraction of holocellulose from fibres was carried out by treating with 17.5% NaOH solution at 20 °C for 45 min and subsequently washed with 10% acetic acid. Later the fibres were treated with 0.8% acetic acid and 0.7% nitric acid in the ratio 15:1 at 120 °C for 15 min. The mixture was cooled, filtered and washed sequentially with 95% ethanol and distilled water. The resulting cellulose fibres were oven dried at 105 °C until consistent weight was achieved.

Characterization

Scanning electron microscopy (SEM)

The morphological structure of gold-sputtered P. pinnata seed hull fibres and isolated cellulose fibres were observed under SEM (JSM-6380LA, JEOL, EVISA). The micrographs were recorded at acceleration voltage of 5–8 kV.

Dynamic light scattering (DLS)

The fibre dimension of aqueous dispersed isolated cellulose fibre (distilled water) was measured by the dynamic light scattering instrument (DLS, nanoparticle analyser, HORIBA Scientific, nano partica SZ-100, Japan).

Fourier transform infrared spectroscopy (FTIR)

Pongamia pinnata seed hull fibres and isolated cellulose fibres mixed with KBr were pressed to form transparent thin pellets. FTIR spectra of the fibres were recorded in the extent of 400–4000/cm with 4/cm resolution using FTIR instrument (Jasco 4200, Jasco analytical instruments, USA).

X-ray analysis (XRD)

XRD measurements for P. pinnata seed hull fibres and isolated cellulose fibres were obtained by X-ray diffractometer (X’Pert3 Powder, PANalytical, The Netherlands) using Cu Kα radiation (1.5406 Å) with Ni filtered at 40 kV, 15 mA. Scattered radiations were recorded in the range of 2θ = 10° − 30° at a scan rate of 4°/min. The Segal method [Eq. (1)] was used to calculate crystallinity index (CI) considering the intensities of \(\left( {2 0 0} \right)\) peak (I 200, 2θ = 22.6°) and the intensity minimum between the \((2 0 0)\) and \((1 1 0)\) peaks (I am, 2θ = 180), where I 200 represents the intensities of crystalline and amorphous material and I am for the amorphous material.

$${\text{CI}} \% = \left( {1 - \frac{{I_{am} }}{{I_{200} }}} \right) \times 100$$
(1)

Thermogravimetric analysis (TGA)

Thermograms for P. pinnata seed hull fibres and isolated cellulose fibres were determined using a thermogravimetric analyser (TGA Q50, TA instruments, USA) at a 10 °C/min heating rate in nitrogen atmosphere.

13C NMR (CP-MAS) spectroscopy

Spectra of P. pinnata seed hull fibres and isolated cellulose fibres were run on solid-state NMR spectrometer (Bruker DSX 300 MHz). 75.46 MHz operating frequency was fixed for 13C nuclei. Fibres were spun at 7.5 kHz spinning rate with filled 5 mm rotor at room temperature.

Results and discussion

Delignification of seed hull using acidified sodium chlorite was compassed, as an initial step in the isolation of cellulose. The alkaline treatment aids in the oxidation of lignin and hemicellulose, solubilizes the residual lignin and hemicellulose resulting in the isolation of cellulose fibres. These cellulose fibres were characterized for their morphological features, thermal stability and also to ensure removal of matrix components such as lignin and hemicellulose.

SEM analysis

The scanning electron microscope images of P. pinnata seed hull fibre after different stages of chemical treatment are as presented in Fig. 1a–c. Dewaxed seed hull fibre presented in Fig. 1a show irregular appearance due to cellulose fibre embedded between waxes and cementing materials such as lignin and hemicellulose (Reddy and Yang 2005; Haafiza et al. 2013). The fibres after sodium chlorite bleaching show cellulose fibres emerging out of the matrix as shown in Fig. 1b. This could be accounted to oxidation and solubilisation of matrix components viz. lignin and hemicellulose. The cementing components—lignin and hemicellulose isolated from the fibres are dissolved by mild alkali treatment (Elanthikkal et al. 2010). As a result, the SEM image of the isolated cellulose fibres as presented in Fig. 1c illustrates individualized single strand of cellulose fibres of diameter 6–8 µm, which in turn is a bundle of cellulose microfibres (Chen et al. 2011) having diameter of 270–370 nm. The cellulose fibres isolated in this work were of smaller diameter compared to that of the other cellulose fibres obtained from different sources such as soybean straw (Reddy and Yang 2009) yielding fibres of diameter 15.6 µm and coconut palm sheath (Maheswari et al. 2012) yielding fibres of 10–15 µm diameter.
Fig. 1
Fig. 1

Scanning electron microscope images a dewaxed Pongamia pinnata seed hull, b sodium chlorite-treated fibres and c isolated cellulose fibres

DLS analysis

The aqueous dispersion of cellulose fibres was analysed by the dynamic light scattering technique in order to find their size distribution. DLS analysis results are summarized in Table 1. The histogram presented in Fig. 2 shows the presence of two distributions, indicating the presence of two dimensions (Kavitha et al. 2013; Srinivas et al. 2012) which is owing to the fibrous structure of cellulose representing both length and diameter. de Carvalho Mendes et al. (2015) also reported such two peaks in DLS histogram of the aqueous dispersion of cellulose fibrous structure. Dimensions determined by DLS epitomise hydrodynamic size (sphere size) having same diffusional coefficient as the fibres being measured (Horiba knowledgebase 2017). The mean hydrodynamic size of isolated cellulose fibres for shorter dimension (diameter) was observed to be 58.4 nm, whereas the longer dimension (length) of the fibres was observed to be 536.3 nm with the standard deviation of 3.3 and 44.1 nm, respectively. The diameter of the fibre obtained by SEM analysis is lesser than that obtained by DLS technique. The sizes of the fibre obtained by SEM and DLS are not comparable, as the diameter obtained by SEM presents the dry fibre size, whereas that obtained by DLS signifies the hydrodynamic diameter in aqueous dispersion. The difference in size estimated by the two methods is generally higher for the nonspherical particles.
Table 1

Particle size distribution values of isolated cellulose fibre

Peak No.

S.P. area ratioa

Mean (nm)

SDb (nm)

Mode (nm)

1

0.20

58.4

3.3

58.9

2

0.80

536.3

44.1

535.0

Total

1.00

441.9

194.3

535.00

aSpecific particle surface area ratio

bStandard deviation

Fig. 2
Fig. 2

DLS analysis spectra of isolated cellulose fibres

Fourier transform infrared spectroscopy (FTIR)

FTIR spectroscopy monitors the functional groups present in the fibres. Figure 3a and b present the spectra obtained for P. pinnata seed hull fibres and isolated cellulose fibres. The band around 3600–3000/cm assigned to stretching vibrations of O–H and C–H is observed in both P. pinnata seed hull fibres and isolated cellulose fibre, indicating the presence of cellulose-related functional groups (Qiao et al. 2016; Shin et al. 2012; Kalita et al. 2015; Sun et al. 2004a, b, c; Kaushik and Singh 2011). Peaks at 2894.63 and 2919.7/cm is generally assigned to C–H stretching vibration in lignin polysaccharide (cellulose and hemicellulose) (Shin et al. 2012; Sun et al. 2004a, b, c, d; Kaushik and Singh 2011; Zhong et al. 2013). Peak at 1735.62/cm is assigned to C=O stretching vibration of carbonyl, acetyl and uronic ester group of the ferulic and p-coumaric acids of lignin and/or xylan component of hemicellulose. The disappearance of these peaks in cellulose fibre spectra, confirms the removal of lignin and hemicellulose (Kalita et al. 2015; Kaushik and Singh 2011; Sun et al. 2004a, c, d; Elanthikkal et al. 2010; Rosa et al. 2012; Oun and Rhim 2016). Peaks at 1646.91 and 1648.84/cm are attributed to O–H bending of absorbed water and are observed in both the spectra; the presence of water could be related to the hydrophilic nature of cellulose component even though the samples analysed were dry (Qiao et al. 2016; Sun et al. 2004b, c; Kaushik and Singh 2011; Zhong et al. 2013; Rosa et al. 2012; Oun and Rhim 2016; Haafiza et al. 2013). Peaks at 1457.92 and 1423.21/cm are usually attributed to aromatic C=C stretch of lignin and the reduction of peak at 1423.21/cm in cellulose fibre spectra indicates the fractional delignification after the treatments (Sun et al. 2004a, b, c, d; Kaushik and Singh 2011; Elanthikkal et al. 2010; Haafiza et al. 2013). Peaks around 1373.07 and 1168.65/cm observed in P. pinnata seed hull fibres are assigned to C–H asymmetric deformation and C–O antisymmetric bridge stretching, respectively (Kalita et al. 2015; Sun et al. 2004a, b, c, d; Kaushik and Singh 2011; Zhong et al. 2013; Rosa et al. 2012). Finally the increase in peak 1033.66/cm, observed in isolated cellulose fibre spectra attributed to –C–O–C– pyranose ring skeletal vibration which indicates an increase in cellulose content (Sun et al. 2004a, b; Elanthikkal et al. 2010).
Fig. 3
Fig. 3

FTIR spectra of a Pongamia pinnata seed hull fibres, b isolated cellulose fibres

13C NMR (CP-MAS) spectroscopy

The 13C NMR spectra of untreated P. pinnata seed hull fibres and isolated cellulose are as shown in Fig. 4a, b. P. pinnata seed hull fibres spectrum in Fig. 4a, illustrates the presence of corresponding signals for the cellulose, hemicellulose and lignin, whereas in the case of isolated cellulose fibre spectrum as shown in Fig. 4b peaks of only cellulose carbon atoms were illustrated. Peaks between 107 and 60 ppm corresponding to six carbon atoms assigned to cellulose molecules are observed in both the spectra. The cellulose carbon atom peak at 107.6 is associated with C1 (Halonen et al. 2013), peaks at 77–67 ppm are assigned to C2, C3 and C5 carbon atoms (Sun et al. 2004a, d), peaks at 91.454 − 84.447 are of C4 (Bhattacharya et al. 2008) and finally 65.305 − 58 is associated with C6 carbon atom (Sun et al. 2004b, c, d). Similar observations were reported by Halonen et al. 2013, where the peaks around 109 − 101 ppm were associated with C1 atom, 80 − 68 ppm to C2, C3 and C5, 91 − 80 ppm to C4 and 68 − 58 ppm to C6 (Bhattacharya et al. 2008). In case of cellulose spectrum, the absence of peaks at 20–33 and 110–140 ppm associated with methylenes in lignin and 58.896 ppm of –OCH3 groups in lignin and hemicellulose, ensures the removal of hemicellulose and lignin, the matrix components (Sun et al. 2004d; Bhattacharya et al. 2008).
Fig. 4
Fig. 4

The 13C NMR spectra of a untreated Pongamia pinnata seed hull fibres, b isolated cellulose fibres

Thus the removal of hemicellulose and lignin from the P. pinnata seed hull fibres are supported by both NMR and FTIR spectral data.

Thermogravimetric analysis (TGA)

The thermograms of untreated P. pinnata seed hull fibres and isolated cellulose fibres as shown in Fig. 5 have onset degradation temperature of 200 and 270 °C, respectively. The major degradation peak at around 250–350 °C observed for isolated cellulose fibre is mainly due to pyrolysis of cellulose and thermal depolymerisation of hemicellulose (Abraham et al. 2011; Li et al. 2015; Chen et al. 2011; Luduena et al. 2011), showing 75% degradation of cellulose. The increase in the decomposition temperature of the isolated cellulose fibres is related to the crystallinity of cellulose due to the removal of lignin and amorphous hemicelluloses (Abe and Yano 2009). Residual presence in both P. pinnata seed hull fibres and isolated cellulose fibres at 800 °C was observed to be 25 and 7%, respectively, which indicates reduction in the presence of carbonaceous materials in the nitrogen atmosphere which is associated with the removal of hemicellulose (Li et al. 2015). Thus the high thermal properties perceived in case of isolated cellulose microfibres may broaden the fields of application of cellulose fibres at temperatures above 200 °C especially for biocomposite processing.
Fig. 5
Fig. 5

Thermogram of untreated Pongamia pinnata seed hull fibres and isolated cellulose fibres

X-ray diffraction (XRD)

X-ray diffractograms of P. pinnata seed hull fibres and isolated cellulose fibres are presented in Fig. 6. Two peaks are observed at 2θ = 16° and 22.6° for both the samples which is the characteristic of crystal polymorphs of cellulose I and cellulose II, respectively (Bondeson et al. 2006; Novo et al. 2015). The peak at 2θ = 16° corresponds to the \((1 1 0)\) and 2θ = 22.6° corresponds to the \((2 0 0)\). The crystallinity index (CI) obtained using Eq. (1) for P. pinnata seed hull fibres and isolated cellulose fibres were 27.2, and 47%, respectively. The crystallinity of the isolated cellulose microfibres was increased by 72.79%. This could be due to the presence of large amount of crystalline cellulose and removal of amorphous hemicellulose and lignin (Rosa et al. 2010) from isolated cellulose fibres by chlorination and alkaline treatment.
Fig. 6
Fig. 6

X-ray diffraction patterns of untreated Pongamia pinnata seed hull and isolated cellulose fibres

Thus, from the above results it can be observed that cellulose microfibres isolated from P. pinnata seed hull exhibited enhanced morphological, thermal and crystalline properties after chlorination and alkaline treatment. Size and increase in crystallinity of the cellulose fibres obtained from different sources and isolation methods are summarized in Table 2. The size of the fibres obtained in the present work is comparable with that obtained from other sources by different isolation methods. However, percentage increase in crystallinity for the fibres isolated from P. pinnata seed hull after chlorination and alkaline treatment is higher than that for the fibres isolated from other sources by chemical treatment methods obtained by other researchers. As observed in Table 2, increase in crystallinity is lower in most of the cases in spite of additional mechanical treatments. Julie et al. (2016) have obtained around 97% increase in crystallinity of the fibres isolated from Arecanut husk fibres. However, they have adopted homogenization, a mechanical process after chemical treatment. Isolation of cellulose microfibres by chlorination and alkaline treatment is economical compared to others, as enormous amount of energy is consumed in the mechanical treatments. The chlorination and alkaline treatment on P. pinnata seed hull resulted in the isolation of crystalline cellulose fibres of 6–8 μm diameter. It is observed that the cellulose fibres isolated from P. pinnata seed hull show higher percentage increase in crystallinity when compared to cellulose fibres obtained from other resources by chemical treatments. Higher crystallinity of cellulose fibres accounts to higher tensile strength of the fibres (Alemdar and Sain 2008), which in turn is expected to enhance the mechanical properties of the cellulose fibre-reinforced composites.
Table 2

Comparison of fibre size and crystallinity index (CI) of cellulose fibre isolated from different sources and isolation treatments

S. no

Source

Size (diameter) of cellulose fibres as observed under SEM

Crystallinity index (CI) isolated cellulose fibre (%)

Increase in crystallinity (%)

Treatment

Reference

1

Resak’s hardwood waste

7–12 (μm)

68.1

37.33

Alkaline and acid hydrolysis

(Abidin et al. 2015)

2

Corn husk

5–8 (nm)

63.3

46.5

Alkaline, TEMPO oxidation, and homogenization

(Du et al. 2016)

3

Arecanut husk fibre

3–5 (nm)

73

97

Alkaline, acid hydrolysis, Bleaching (Chlorination), homogenization

(Julie et al. 2016)

4

Moso bamboo culms

0.5–1 (μm)

65.32

28.87

Microwave liquefaction, bleaching (Chlorination), Alkaline, homogenization and ultrasonication

(Xie et al. 2016)

5

Gigantochloa scortechinii bamboo culms

5.29–10.94 (nm)

65.32

36.33

Acid hydrolysis, homogenization, sonication

(Saurabh et al. 2016)

6

Sugarcane bagasse

<20 (μm)

–

–

Steam explosion, enzymatic treatment

(Saelee et al. 2014)

7

Mengkuang leaves

5–80 (μm)

69.5

26.13

Alkaline Bleaching (Chlorination), sonication

(Sheltami et al. 2012)

8

Rice husk

7 (μm)

59

26.06

Alkali, bleaching

(Johar et al. 2012)

9

Coconut palm leaf sheath

10–15 (μm)

47.7

12.7

Chlorination and alkaline

(Maheswari et al. 2012)

10

Pongamia pinnata seed hull

6–8 (μm)

47

72.79

Chlorination and alkaline

Present work

Conclusion

Cellulose fibres were isolated from P. pinnata seed hull by sequential chlorination and alkaline process and the resultant microfibres were characterized by SEM, DLS, FTIR, NMR, TGA and XRD analyses. Cellulose microfibres were in diameter ranging from 6 to 8 μm and mean hydrodynamic diameter of 58.4 nm. NMR and FTIR analyses confirmed the removal of hemicellulose and lignin. Crystallinity of the fibres was increased by 72.79% after the treatment with CI of 47% for the isolated cellulose fibres. Thermal behaviour of the fibres had improved as evidenced by an increase of degradation temperature by 70 °C. Most potential observation to be considered was the degradation temperature of the isolated cellulose fibres being higher than 200 °C, which could broaden its application potential in the fields of biocomposite processing. A notable increase in crystallinity and the dimension similar to the cellulose fibres isolated from other resources by various chemical treatments was a significant feature of the resource and the isolation method adopted in the present study. Thus the present work substantiates the success of the sequential chlorination and alkaline extraction process solely contributing to obtain smaller diameter and crystalline cellulose microfibres from P. pinnata seed hull. These biofibres have potential application as filler, embroiling in the process of biodegradable composites to enhance their properties.

Declarations

Authors’ contributions

PM, GS and KVS are the primary contributors, as this work is the result of Ph.D. work. All authors read and approved the final manuscript.

Acknowledgements

Authors gratefully acknowledge University of Agricultural Sciences, Bengaluru, India for providing Pongamia pinnata seed hull.

Competing interests

The authors declare that they have no competing interests.

Consent for publication

All authors declare that consent submitted for the final accepted version of the manuscript to be considered for publication in Bioresources and Bioprocessing journal.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors’ Affiliations

(1)
Department of Chemical Engineering, National Institute of Technology Karnataka, Surathkal, India

References

  1. Abe K, Yano H (2009) Comparison of the characteristics of cellulose microfibril aggregates of wood, rice straw and potato tuber. Cellulose 16:1017–1023View ArticleGoogle Scholar
  2. Abidin NAMZ, Aziz FA, Radiman S, Ismail A, Yunus WMZW, Nor NM, Sohaimi RM, Sulaiman AZ, Halim NA, Darius DDI (2015) Isolation of microfibrillated cellulose (MFC) from local hardwood waste, Resak (Vatica spp.). Mater Sci Forum 846:679–682. doi:10.4028/www.scientific.net/MSF.846.679 View ArticleGoogle Scholar
  3. Abraham E, Deepa B, Pothan LA, Jacob M, Thomas S, Cvelbar U, Anandjiwala R (2011) Extraction of nanocellulose fibrils from lignocellulosic fibers: a novel approach. Carbohydr Polym 86(4):1468–1475View ArticleGoogle Scholar
  4. Alemdar A, Sain M (2008) Isolation and characterization of nanofibers from agricultural residues—wheat straw and soyhulls. Bioresour Technol 99(6):1664–1671View ArticleGoogle Scholar
  5. Bhattacharya D, Germinario LT, Winter WT (2008) Isolation, preparation and characterization of cellulose microfibers obtained from bagasse. Carbohydr Polym 73:371–377View ArticleGoogle Scholar
  6. Bondeson D, Mathew A, Oksman K (2006) Optimization of the isolation of nanocrystals from microcrystalline cellulose by acid hydrolysis. Cellulose 13:171–180View ArticleGoogle Scholar
  7. Chen Y, Liu C, Chang PR, Anderson DP, Huneault MA (2009) Pea starch-based composite films with pea hull fibers and pea hull fiber-derived nanowhiskers. Polym Eng Sci 49(2):369–378. doi:10.1002/pen.21290 View ArticleGoogle Scholar
  8. Chen W, Yu H, Liu Y, Chen P, Zhang M, Hai Y (2011) Individualization of cellulose nanofibers from wood using high-intensity ultrasonication combined with chemical pretreatments. Carbohydr Polym 83:1804–1811View ArticleGoogle Scholar
  9. Cherian BM, Leão AL, de Souza SF, Thomas S, Pothan LA, Kottaisamy M (2010) Isolation of nanocellulose from pineapple leaf fibers by steam explosion. Carbohydr Polym 81:720–725View ArticleGoogle Scholar
  10. de Carvalho Mendes CA, Ferreira MS, Furtado CRG, de Sousa AMF (2015) Isolation and characterization of nanocrystalline cellulose from corn husk. Mater Lett 148:26–29View ArticleGoogle Scholar
  11. Demirbas A (2009) Heavy metal adsorption onto agro based waste materials: a review. J Hazard Mater 157(2–3):220–229Google Scholar
  12. Dinand E, Chanzy H, Vignon RM (1999) Suspensions of cellulose microfibrils from sugar beet pulp. Food Hydrocolloid 13(3):275–283View ArticleGoogle Scholar
  13. Du C, Li H, Li B, Liu M, Zhan H (2016) Characteristics and properties of cellulose nanofibers prepared by TEMPO oxidation of corn husk. BioResources 11(2):5276–5284Google Scholar
  14. Dufresne A, Cavaille JY, Vignon MR (1997) Mechanical behavior of sheets prepared from sugar beet cellulose microfibrils. J Appl Polym Sci 64:1185–1194View ArticleGoogle Scholar
  15. Elanthikkal S, Gopalakrishnapanicker U, Varghese S, Guthrie JT (2010) Cellulose microfibers produced from banana plant wastes: isolation and characterization. Carbohydr Polym 80:852–859View ArticleGoogle Scholar
  16. Espino E, Cakir M, Domenek S, Román-Gutiérrez AD, Belgacem N, Bras J (2014) Isolation and characterization of cellulose nanocrystals from industrial by-products of Agave tequilana and barley. Ind Crops Prod 62:552–559. doi:10.1016/j.indcrop.2014.09.017 View ArticleGoogle Scholar
  17. Fiore V, Scalici T, Valenza A (2014) Characterization of a new natural fiber from Arundo donax L. as potential reinforcement of polymer composites. Carbohydr Polym 106:77–83View ArticleGoogle Scholar
  18. Haafiza MKM, Eichhornc SJ, Hassana A, Jawaid M (2013) Isolation and characterization of microcrystalline cellulose from oil palm biomass residue. Carbohydr Polym 93(2):628–634View ArticleGoogle Scholar
  19. Halonen H, Larsson PT, Iversen T (2013) Mercerized cellulose biocomposites: a study of influence of mercerization on cellulose supramolecular structure, water retention value and tensile properties. Cellulose 20:57–65View ArticleGoogle Scholar
  20. Horiba knowledgebase (2017) Horiba Instruments, Inc. https://www.horiba.com/fileadmin/uploads/Scientific/eMag/PSA/Guidebook/pdf/PSA_Guidebook.pdf. Accessed 05 Nov 2016
  21. Hou X, Sun F, Zhang L, Luo J, Lu D, Yang Y (2014) Chemical-free extraction of cotton stalk fibers by steam flash explosion. BioResources 9(4):6950–6967View ArticleGoogle Scholar
  22. Hubbe MA, Rojas OJ, Lucia LA, Sain M (2008) Cellulosic nanocomposites: a review. BioResources 3:929–980Google Scholar
  23. Johar N, Ahmad I, Dufresnec A (2012) Extraction, preparation and characterization of cellulose fibres and nanocrystals from rice husk. Ind Crops Prod 37:93–99View ArticleGoogle Scholar
  24. Julie CCS, George N, Narayanankutty SK (2016) Isolation and characterization of cellulose nanofibrils from arecanut husk fibre. Carbohydr Polym. doi:10.1016/j.carbpol.2016.01.015 Google Scholar
  25. Kalita E, Nath BK, Deb P, Agan F, Islam MR, Saikia K (2015) High quality fluorescent cellulose nanofibers from endemic rice husk: isolation and characterization. Carbohydr Polym 122:308–313View ArticleGoogle Scholar
  26. Kaushik A, Singh M (2011) Isolation and characterization of cellulose nanofibrils from wheat straw using steam explosion coupled with high shear homogenization. Carbohydr Res 346(1):76–85.View ArticleGoogle Scholar
  27. Kavitha B, Kumar KS, Narsimlu N (2013) Synthesis and characterization of polyaniline nano-fibers. Indian J Pure Appl Phys 51:207–209Google Scholar
  28. Li R, Fei J, Cai Y, Li Y, Fengand J, Yao J (2009) Cellulose whiskers extracted from mulberry: a novel biomass production. Carbohydr Polym 76(1):94–99View ArticleGoogle Scholar
  29. Li W, Zhang Y, Li J, Zhou Y, Li R, Zhou W (2015) Characterization of cellulose from banana pseudo-stem by heterogeneous liquefaction. Carbohydr Polym 132:513–519View ArticleGoogle Scholar
  30. Luduena L, Fasce D, Alverez VA, Stefani PM (2011) Nanocellulose from rice husk following alkaline treatment to remove silica. BioResources 6(2):1440–1453Google Scholar
  31. Maheswari UC, Obi Reddy K, Muzenda E, Guduri BR, Varada Rajulu A (2012) Extraction and characterization of cellulose microfibrils from agricultural residue—Cocos nucifera L. Biomass Bioenerg 46: 555–563. doi 10.1016/j.biombioe.2012.06.039. http://www.sciencedirect.com/science/journal/0961953446:555–563
  32. Mandal A, Chakrabarty D (2011) Isolation of nanocellulose from waste sugarcane bagasse (SCB) and its characterization. Carbohydr Polym 86:1291–1299View ArticleGoogle Scholar
  33. Mangal R, Saxena NS, Sreekala MS, Thomas S, Singh K (2003) Thermal properties of pineapple leaf fiber reinforced composites. Mater Sci Eng 339:281–285View ArticleGoogle Scholar
  34. Moran JI, Alvarez VA, Cyras VP (2008) Extraction of cellulose and preparation of nanocellulose from sisal fibers. Cellulose 15:149–159. doi:10.1007/s10570-007-9145-9 View ArticleGoogle Scholar
  35. Motaung TE, Mtibe A (2015) Alkali treatment and cellulose nanowhiskers extracted from maize stalk residues. Mater Sci Appl 6:1022–1032. doi:10.4236/msa.2015.611102 Google Scholar
  36. Nadeem R, Ansari TM, Akhtar K, Khalid AM (2009) Pb (II) sorption by pyrolysed Pongamia pinnata pods carbon (PPPC). Chem Eng J 152:54–63. doi:10.1016/j.cej.2009.03.030 View ArticleGoogle Scholar
  37. Novo LP, Bras J, García A, Belgacem N, Curvelo AA (2015) Subcritical water: a method for green production of cellulose nanocrystals. ACS Sustain Chem Eng 3:2839–2846View ArticleGoogle Scholar
  38. Oun AA, Rhim JW (2016) Characterization of nanocelluloses isolated from Ushar (Calotropis procera) seed fiber: effect of isolation method. Mater Lett 168:146–150View ArticleGoogle Scholar
  39. Qiao C, Chen G, Zhang J, Yao J (2016) Structure and rheological properties of cellulose nanocrystals suspension. Food Hydrocolloid 55:19–25View ArticleGoogle Scholar
  40. Reddy N, Yang Y (2005) Biofibers from agricultural byproducts for industrial applications. Trends Biotechnol 23(1):22–27View ArticleGoogle Scholar
  41. Reddy N, Yang Y (2009) Natural cellulose fibers from soybean straw. Bioresour Technol 100:3593–3598View ArticleGoogle Scholar
  42. Rosa MF, Medeiros ES, Malmonge JA, Gregorski KS, Wood DF, Mattoso LHC (2010) Cellulose nanowhiskers from coconut husk fibers: effect of preparation conditions on their thermal and morphological behavior. Carbohydr Polym 81(1):83–92View ArticleGoogle Scholar
  43. Rosa SM, Rehman N, de Miranda MIG, Nachtigall SM, Bica CI (2012) Chlorine-free extraction of cellulose from rice husk and whisker isolation. Carbohydr Polym 87:1131–1138View ArticleGoogle Scholar
  44. Saelee K, Yingkamhaeng N, Nimchua T, Sukyai P (2014) Extraction and characterization of cellulose from sugarcane bagasse by using environmental friendly method. In: Proceedings of The 26th Annual Meeting of the Thai Society for Biotechnology and International Conference, Mae Fah Lunag University (School of Science), Thailand, 26–29 November 2014Google Scholar
  45. Saurabh CK, Mustapha A, Masri MM, Owolabi AF, Syakir MI, Dungani R, Paridah MT, Jawaid M, Khalil HPSA (2016) Isolation and characterization of cellulose nanofibers from Gigantochloa scortechinii as a reinforcement material. J Nanomater. doi:10.1155/2016/4024527 Google Scholar
  46. Sheltami RM, Abdullaha I, Ahmada I, Dufresnec A, Kargarzadeha H (2012) Extraction of cellulose nanocrystals from mengkuang leaves (Pandanus tectorius). Carbohydr Polym 88:772–779View ArticleGoogle Scholar
  47. Shin HK, Jeun JP, Kim HB, Kang PH (2012) Isolation of cellulose fibers from kenaf using electron beam. Radiat Phys Chem 81:936–940View ArticleGoogle Scholar
  48. Shwetha KC, Nagarajappa DP, Mamatha M (2014) Removal of copper from simulated wastewater using Pongamia pinnata seed shell as adsorbent. Int J Eng Res Appl 4(6):271–282Google Scholar
  49. Srinivas CH, Srinivasu D, Kavitha B, Narsimlu N, Siva Kumar K (2012) Synthesis and characterization of nano size conducting polyaniline. IOSR J Appl Phys 1(5):12–15View ArticleGoogle Scholar
  50. Sun JX, Sun XF, Sun RC, Su YQ (2004a) Fractional extraction and structural characterization of sugarcane bagasse hemicelluloses. Carbohydr Polym 56:195–204View ArticleGoogle Scholar
  51. Sun JX, Sun XF, Zhao H, Sun RC (2004b) Isolation and characterization of cellulose from sugarcane bagasse. Polym Degrad Stabil 84:331–339View ArticleGoogle Scholar
  52. Sun XF, Sun RC, Su Y, Sun JX (2004c) Comparative study of crude and purified cellulose from wheat straw. J Agric Food Chem 52(4):839–847View ArticleGoogle Scholar
  53. Sun XF, Sun RC, Fowler P, Baird MS (2004d) Isolation and characterisation of cellulose obtained by a two-stage treatment with organosolv and cyanamide activated hydrogen peroxide from wheat straw. Carbohydr Polym 55:379–391View ArticleGoogle Scholar
  54. Xie J, Hse CY, De Hoop CF, Hu T, Qi J, Shupe TF (2016) Isolation and characterization of cellulose nanofibers from bamboo using microwave liquefaction combined with chemical treatment and ultrasonication. Carbohydr Polym 151:725–734View ArticleGoogle Scholar
  55. Yadav PP, Ahmed G, Maurya R (2004) Furanoflavonoids from Pongamia pinnata fruit. Phytochemistry 65(4):439–443View ArticleGoogle Scholar
  56. Zhong C, Wang C, Huang F, Jia H, Wei P (2013) Wheat straw cellulose dissolution and isolation by tetra-n-butylammonium hydroxide. Carbohydr Polym 94:38–45View ArticleGoogle Scholar
  57. Zuluaga R, Putaux JL, Cruz J, Velez J, Mondragon I, Ganan P (2009) Cellulose microfibrils from banana rachis: effect of alkaline treatments on structural and morphological features. Carbohydr Polym 76:51–59View ArticleGoogle Scholar

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