Skip to main content
Download PDF

Ultrasonic intensification as a tool for enhanced microbial biofuel yields

Biotechnology for Biofuels volume 8, Article number: 140 (2015) Cite this article

Abstract

Ultrasonication has recently received attention as a novel bioprocessing tool for process intensification in many areas of downstream processing. Ultrasonic intensification (periodic ultrasonic treatment during the fermentation process) can result in a more effective homogenization of biomass and faster energy and mass transfer to biomass over short time periods which can result in enhanced microbial growth. Ultrasonic intensification can allow the rapid selective extraction of specific biomass components and can enhance product yields which can be of economic benefit. This review focuses on the role of ultrasonication in the extraction and yield enhancement of compounds from various microbial sources, specifically algal and cyanobacterial biomass with a focus on the production of biofuels. The operating principles associated with the process of ultrasonication and the influence of various operating conditions including ultrasonic frequency, power intensity, ultrasonic duration, reactor designs and kinetics applied for ultrasonic intensification are also described.

Background

The utilisation of algae and cyanobacteria as cell factories for the production of biofuels has recently received considerable attention. Their ability to grow photoautotrophically and provide a renewable and sustainable biofuel source whilst also providing additional opportunities for co-product biorefinery makes them intriguing candidates [1]. There have been many reports that algal biofuel production is technically if as yet not economically viable [2–5]. Therefore, any technique that improves the efficiency of biofuel production is likely to have a significant effect on the viability of these production processes.

Algae and cyanobacteria can be metabolically engineered to produce biofuels such as ethanol [6, 7], biogas [8], hydrogen [9, 10], free fatty acids for transesterification to biodiesel [11, 12] and butanol [13]. Furthermore, there are several ways to convert algal or cyanobacterial biomass to fuel energy such as biochemical conversion, chemical treatment, direct combustion and thermochemical conversion [14]. Lipids can be extracted and transesterified into biodiesel while starch, glycogen and carbohydrate polymers, can be further hydrolyzed and converted into bio-ethanol [14]. Indeed following lipid extraction from Chlorococum sp. the residual biomass, used as a feedstock, has been shown to produce significant amounts of bioethanol upon fermentation with concentrations of 3.83 g L−1 achievable [15, 16]. In this case, integration of an ultrasonication pre-treatment, known to enhance lipid extraction yields and biogas production can also be quite beneficial in disrupting the spent biomass [8, 17, 18]. This review aims to examine the beneficial aspects of ultrasonic intensification of lipid extraction, fermentation and production of biofuels.

Ultrasonic process intensification: concept introduction

Ultrasonication is a branch of acoustics that can be applied to solids, liquids and gases at frequencies above the human hearing range [19]. The frequencies applied for processes are shown in Fig. 1. Particle agitation in a liquid culture can be achieved by applying acoustic energy with frequencies ranging from 10 kHz to 20 MHz using ultrasonic probes, an ultrasonic bath, a flat plate or a tube type ultrasonicator [20]. The process operates by converting electrical energy into physical vibration which directly influences the medium it is applied to by imparting high energy to the medium via cavitation [21]. During the vibration process, the microbubbles present in the form of nuclei are increased in size to a maximum of about 4–300 mm in diameter [22], and can be either stable or transient. In the case of low ultrasonic intensity, the radii of microbubbles periodically and repetitively expand and shrink causing radial oscillation within several acoustic cycles. At the point when acoustic energy has sufficient intensity, some microbubbles become unstable and when the resonant frequency of bubbles exceeds that of the ultrasonic field, the bubbles collapse within a few nanoseconds at the solid/solvent interface (>200 mm) [21] which produces microjets with a velocity >100 m s−1 and shock waves of approximately 103 MPa towards the solid surface of the substance in solution. This causes cavitation of the substance in the liquid medium, with the violent movement of fluid towards or away from the cavitational microbubbles defined as micro-convection [21]. The convection in the ultrasonic system has two components, i.e., microturbulence or micro-convection and shock waves [23]. The authors have stated that micro-convection is a continuous oscillatory motion of liquid medium induced by radial movement of cavitation bubble and governs the growth of the nuclei while shock waves are discrete, high pressure amplitude waves emitted by the bubble which increase the nucleation rate [23]. This micro cavitation influences the transport of fluids and solid particles within the medium and results in forces that can cause emulsification or dispersion, while the strong shockwaves and microjets generate extremely strong shear forces over those of conventional mechanical methods, and are able to scatter liquid into tiny droplets or crush solid particles into fine powders [21].

Fig. 1
figure 1

Ultrasonic frequency scale for various fields of technology

Full size image

Ultrasonication which enhances biochemical reactions is termed ultrasonic process intensification [22] and can be differentiated into low and high intensity applications. In several biotechnology processes, both high and low intensity ultrasonic waves have been utilised depending on the objective of the sonication process [24]. Low intensity ultrasonic (<1 W cm−2 and between 1 and 10 MHz) intensification can be considered non-destructive as it sends ultrasonic waves through a liquid medium without causing any permanent physical or chemical change in the medium or microorganisms within. The low intensity ultrasonication can also be defined in terms of acoustic pressure amplitude. When the acoustic pressure amplitude is less than the static pressure in the medium, the bubbles undergo stable, small amplitude radial motion called “stable cavitation”. The bubble motion turns transient when the acoustic pressure amplitude exceeds the static pressure in the medium. The microorganisms in the medium respond to the low energy only during the time of exposure to the ultrasonic waves and return to an equilibrium state when the ultrasonic source is removed [24]. On the other hand, high intensity ultrasonic intensification or low frequency ultrasonication (10–1000 W cm−2 and 10–100 kHz) which generates high pressure in the medium can disrupt microbial cellular structures [24]. This can cause the lysis of microbial cells or the formation of free radicals in chemical degradation reactions [25]. Thus, low intensity ultrasonic intensification is quite distinct and transitory when applied periodically. The type of cavitation generated during ultrasonic process intensification depends on several parameters, including amplitude, pressure, temperature, viscosity and concentration of the medium [26].

Physical mechanism of ultrasound assisted processes for biofuels production

Ultrasound causes both physical and chemical changes through the process of cavitation. Chemically, highly reactive radicals can be generated from the dissociation of the entrapped vapour molecules in a cavitation bubble [27]. Physically, cavitation can cause intense convection in a bulk medium leading to microturbulence (an intense oscillating motion of liquid with low to moderate velocities) and shock waves (high pressurised waves emitted by the bubble, with amplitudes as high as 30–50 bar). During lipid extraction from biomass, the physical effects of ultrasonication can significantly enhance the lipid yield. Microturbulence can lead to a more efficient mixing of the biomass and solvent (without induction of shear stress), while shock waves can cause rupture of the cell wall [28]. Ultrasound can also generate intense local turbulence in the medium, pushing the extracted lipids away from the surface of the microbial cells, and thus, maintaining a constant concentration gradient for continuous diffusion of lipids from the cells [27].

The physical effects of ultrasonication can also enhance the transesterification process during biodiesel production [29–32]. Ultrasonication generates an enormous interfacial area between the oil and alcohol due to microturbulence leading to the formation of fine emulsions [33]. Kalva et al. [29] investigated the mechanism of the enhancement of transesterification by discriminating the physical and chemical effects of ultrasound. This was analysed by transesterifying soybean oil using methanol and sodium hydroxide as the base catalyst with ultrasound frequency set at 20 kHz and power output of 100 W. It was reported that the enhancement of transesterification was due to the physical effect rather than a chemical effect of ultrasound, i.e., production of radical species and acceleration of the reaction by these species [34–36]. In addition it was found that the transesterification yield increased when the alcohol to oil molar ratio was increased. This was due to the effect of low intensity microturbulence generated by the cavitation bubbles in the oil, which restricted the dispersion of oil in methanol at high alcohol to oil molar ratios [29].

Ultrasound may also enhance enzymatic reactions within the cell. Khanna et al. [30] reported that the nature of convection generated by ultrasound at raised static or ambient pressure in the medium provided oscillatory motion of liquid, accompanied by medium amplitude movement by the acoustic or shock waves emitted by the bubbles. This was concluded by investigating the mechanism of enhancement of glycerol conversion into 1,3-propanediol and ethanol using Clostridium pasteurianum MTCC 116 with the aid of ultrasonication. Mild shock waves of ultrasonication were shown to cause rapid movement of microbial cells in the fermentation broth. Inter-collisions and/or collisions with the walls of the test tube lead to the acceleration of enzymatic reactions inside the cells [30]. In addition, it was reported that the high velocity micro-streaming caused desorption of the CO2 produced in the metabolic pathway and thus favoured a faster diffusion of glycerol into the microbial cells. As a result, the overall enzymatic reaction system is expected to be substrate saturated with enzyme being the limiting reactant. During ethanol formation in this system the Michaelis constant (Km) was found to decrease with ultrasonication indicating a rise in the formation of enzyme–substrate complex. Hence, ethanol yields were found to increase from 0.024 to 0.04 mol mol−1 (i.e., 83 % increase) and a decrease in substrate inhibition (KI) contributed to the enhancement of 1,3-propanediol yields from 0.017 to 0.021 mol mol−1 (30.89 % increase) [30, 36].

Ultrasonication has also been shown to enhance ethanol fermentation processes by increasing transport through the cell membrane and increasing enzymatic hydrolysis with strong micro-convection [37–40]. Recent studies have found that it could enhance the utilisation of substrate for cell growth, reduce the inhibition by substrate and decrease the specific death rate of cells leading to an increase of ethanol yield from 0.149 to 0.166 g g−1 of raw biomass [41].

Ultrasonic intensification applicability within extraction processes

Traditionally, solvent based processes are utilised for extraction of many bioactive compounds [42, 43]. However, standard extraction mechanisms such as cell maceration and soxhlet extraction have limitations such as high solvent consumption, large operating cost and extended operation times which frequently result in lower yields [44]. The application of ultrasonic intensification can allow higher yields to be generated in a shorter time, with lower energy input [45] and without adding additional reagents to the extraction. In addition, the effects of increasing temperature on the extraction components can be avoided. Such modified extraction techniques have been developed recently for the extraction of macromolecules such as polysaccharides, proteins, terpenoids, flavonoids, carotenoids and phenolic compounds [46–49]. The ultrasonic intensification process can be effectively used to improve the extraction rate by increasing the mass transfer due to the formation of microcavities leading to higher growth and product yields. Ying et al. [49] reported that ultrasonic-based extraction is associated with two main physical phenomena, acoustic cavitation and diffusion through the cell wall. The conditions associated with cavitation, an increase in temperature and pressures up to 100 MPa, produce very high shear energy waves and turbulence in the cavitation zone. The combination of these factors (pressure, heat and turbulence) is used to accelerate mass transfer in the extraction process. Ultrasonic intensification also exerts a mechanical effect which leads to enhanced diffusion of solvents into the cell wall. In pure liquids, the microbubbles retain their spherical shape during the collapse, as their surroundings are uniform [50]. However, when the microbubbles collapse near a solid surface it occurs asymmetrically and produces shock waves toward the cell wall. These waves have a strong impact on the cell surface; therefore, enhance the solvent penetration into the cell. Another effect caused by the ultrasound wave is that it can facilitate the swelling and hydration of biomass and so cause an enlargement of pores in the cell wall which can improve diffusion processes and therefore enhance mass transfer which can enhance extraction yield [50]. Hence ultrasonic intensification can provide high extraction efficiency in a short time with less solvent consumption over other extraction techniques [51]. As an example, Rocco et al. [52] reported a 73 % increase in recovery of polychlorinated biphenyls from biomass generated through treatment of household wastewater with 30 min of ultrasonic intensification during the extraction process.

Benefits of ultrasonic intensification for lipids extraction in biofuel production systems

Algae and cyanobacteria are of interest as a renewable energy source for biofuel largely because of their autotrophic lifestyle and their potential to produce a range of additional products which can aid the economics of biofuel production [53]. For lipid production in particular, lipid extraction is required and cell disruption is necessary for lipid recovery [54]. Though several disruption methods including compression, high pressure homogenization, autoclaving, bead mill treatment, microwave treatment and magnetic stirring have been employed, both at laboratory and pilot scale, the energy requirement for those techniques is typically higher than the energy of combustion of lipid extracted [55]. See Table 1 for a comparison of these extraction methodologies.

Table 1 Comparison of various algal lipid extraction methods
Full size table

Ultrasonic intensification has been shown to improve lipid extraction via cell disruption with more favourable economics than other disruption methods [60], enhancing the extraction of lipids by 50–500 % compared to traditional methods, with a 10 fold reduction in extraction time [61]. Suganya and Renganathan [62] investigated lipid extraction from the green algae Ulva lactuca using ultrasonication and found that the yield of lipid (8.49 %), was maximised with a 6 min ultrasonic pre-treatment. Lipid extraction with Synechocystis aquatilis also demonstrated that ultrasonication resulted in higher yields of lipid, with 21.30 % extracted compared to grinding (18.74 %), osmotic shock (14.55 %) and non-disruptive methods (10.17 %) [63]. For more examples, see Table 2 detailing the reaction conditions and lipid yields recovered from various microalgae sources using ultrasonic intensification.

Table 2 Reaction conditions for ultrasonic intensification on extraction of lipids from various microalgae
Full size table

Algal cell walls are typically tri-layered rigid structures with high tensile strength [66], hence the release of intra lipids can be blocked. Homogenization can affect the outer cell walls with shearing force but not the interior of the cell. Extraction of lipids from cells may occur by either diffusion of lipids across the cell wall, if the algal biomass is suspended in the solvent with higher selectivity and solubility (or large partition coefficient) for lipids or disruption of the cell wall with release of cell contents in the solvent. The diffusive mechanism is less efficient due to slow diffusion of lipid across the cell wall while disruptive mechanism results in faster extraction with high yield as it causes direct release of lipid due to the rupture of cell wall [27]. Sonication can interfere with the cell interior via shock waves produced by imploding cavitation bubbles [67]. Thus, ultrasonic intensification can be an advantageous addition for lipid extraction processes involving algae. Park et al. [68] investigated the effect of homogenisation and ultrasonication in combination on lipid extraction from Chlorella vulgaris. The initial fatty acid content of C. vulgaris was 360 mg g−1 cell. Lipid recovery was found to increase when both techniques were combined compared with the use of either alone. The results of the combined processes showed 100.5, 123.9, and 152.0 mg lipid g−1 cell recovered for the 20, 40, and 60 min reaction times used, respectively. The yields were 5.3-fold, 6.6-fold, and 8.1-fold higher, respectively, than that of the control (single treatment by sonication or single treatment via homogenization). In this system [68], microalgal suspensions were allowed to circulate continuously between the homogenizer and ultrasonicator and it was concluded that cell walls damaged slightly by homogenization would be effectively disrupted by the subsequent ultrasonication-induced cavitation bubbles resulting in a superior extraction process [68]. When the cell concentration was increased to 40 g.L−1, the lipid recovery yield for 1 h of sonication-assisted homogenization was increased to a very high 281.3 mg lipid g−1 cell using the chloroform–methanol solvent. This system’s treatment of high cell concentrations makes it possible to enhance lipid recovery capacity based on unit time [68].

In addition to enhancing the yield of lipids extracted from cells, low frequency ultrasonic intensification can also enhance emulsion generation with immiscible liquids which are commonly used during biodiesel production. This was confirmed via an alkali catalysed biodiesel production process performed by Ji et al. [69] which found that a 100 % emulsion could be achieved within 20 min with the utilisation of a jacketed ultrasonic reactor [69]. Moreover, the beneficial effects of ultrasonication have been reported for the transesterification of triglycerides with methanol. Reaction times could be reduced, as could the catalyst requirement, the energy consumption and the alcohol to oil molar ratio [70, 71]. Because of these beneficial effects on extraction and transesterification, these processes are frequently carried out concurrently. When both lipid extraction and transesterification are carried out simultaneously, it is referred to as in situ transesterification [72]. In situ transesterification processes using microbial biomass for biodiesel production can greatly enhance the biodiesel yield, as described in Table 3.

Table 3 Effect of ultrasonic intensification on in situ transesterification of various microbial biomasses
Full size table

Ultrasonic intensification, utility in fermentation processes

High and low intensity ultrasonic intensification methods are utilised to enhance industrial fermentation processes including those involving the production of biofuels [73]. Ultrasonic intensification has been used as a pre-treatment prior to the fermentation of algal biomass for the production of bioethanol [74]. Upon ultrasonication, significant changes in the physicochemical properties of the algal cell enhance the bio-accessibility of carbohydrate substrates in the fermentation media, increasing their availability for microbial fermentation. Sonication pre-treatments (frequency 40 kHz; power output 2.2 kW) for 15 min or longer on Scenedesmus obliquus YSW15 resulted in increasing the concentration of dissolved carbohydrate. This increased availability resulted in enhanced ethanol production by up to 5.6 g L−1. In addition, sonication can decrease algal surface hydrophobicity and increase the electrostatic repulsion among the algal debris dispersed in the aqueous solution. This can provide more facile access to the treated algal biomass, enhancing the assimilation of algal carbohydrates by anaerobic bacteria isolated from seed sludge of municipal wastewater [74]. Low intensity ultrasonication has also been found to promote protein production and increase cell concentration through the enhancement of cell membrane porosity [75]. For instance, upon ultrasonication, Pseudomonas aeruginosa membrane porosity has been shown to increase, resulting in the enhanced uptake of 16-doxylstearic acid (a hydrophobic antibiotic) through the cell membrane [76]. In general, such increases in cell permeability lead to enhanced diffusion rates and thereby increase the overall cell productivity and growth rate [77]. In the case of E. coli, the average cell count was almost doubled by the inclusion of an ultrasonication treatment [8.5 × 107 CFU ml−1 versus 4.8 × 107 CFU ml−1] [77]. Indeed the lag time (λ) for non-ultrasonicated biomass during H2 production from the microalgae Scenedesmus obliquus YSW15 was higher than that for the ultrasonicated biomass. λ derived from the Gompertz equation indicated that the non-ultrasonicated biomass had an average 9 h lag phase, which was reduced to 6 h after ultrasonication (45 kHz with power output 2.2 kW). The shorter lag phase in ultrasonicated biomass fermentation was reported due to the easy accessibility of dissolved carbohydrate to fermentative bacteria compared with the non-sonicated control [78]. The maximum H2 production was obtained from 60 min ultrasonicated biomass (2419 mL L−1) whereas the non-sonicated control and short-term ultrasonicated biomass for 5 min produced 1393 and 1370 mL L−1, respectively [78]. Ultrasound (at 20 kHz) has also been demonstrated to induce protein production, specifically thrombinase (a fibrinolytic enzyme) production from marine actinomycetes, likely due to increased mass transfer rates leading to increased nutrient uptake [79]. In conventional bioreactors, microbial cells suspended in the fermentation broth are invariably surrounded by a stagnant film of liquid [80]. This film can hinder the mass transfer of nutrients and products and can be a rate controlling factor [80]. In an ultrasonicated bioreactor, the pulsation of microbubbles of gas in the fluid generates micro-streaming [81] which can minimise the fluid boundary layer around cells located close to the bubbles [82], thus enhancing mass transfer. Ultrasonic intensification can also increase the rates of gas–liquid oxygen transfer, removal of carbon dioxide and dissolution of suspended solids. This can increase the supply of low solubility substrates and, indirectly, enhance microbial cell productivity [83]. Consequently, there are many benefits to the inclusion of a sonication treatment within fermentation processes. Ultrasonic contribution to the enhancement of fermentation yield is given in Table 4.

Table 4 Effect of ultrasonic intensification on fermentative bioenergy
Full size table

The length of ultrasonic treatment has a significant effect on product yields. In a study using c with both continuous and intermittent ultrasonication during fermentation it was found that the ethanol concentration was increased up to 30 g L−1 with intermittent ultrasonic intensification, whereas with continuous ultrasound intensification, no increase was observed [89]. Similarly intermittent ultrasonic intensification (in fermentation broth was found to enhance the production of a fibrinolytic enzyme from Bacillus sphaericus MTCC 3672 by 1.82 fold compared to non-sonicated broth [90]. The intensity of ultrasonication can also affect yields obtained. In a study with molasses fermentation by Saccharomyces cerevisiae M30, the effect of ultrasonic intensification on ethanol production was examined. The ultrasonic frequencies were varied as 20, 25 and 30 kHz with a maximum specific ethanol production rate of 1.55 g g−1 h−1 being achieved at 25 kHz [91].

While low to high intensity pulsing can enhance productivities, increased microbial exposure to ultrasonic power causes cells to become fragile and reduces microbial viability due to mechanical stress on the microbial cell [92]. The mechanical shear caused by the ultrasonic waves can disrupt the cell wall at higher duty cycles and ultimately has a lethal effect [93]. Thus, numerous studies have shown that intermittent ultrasonic intensification is more advantageous, increasing product yield while keeping microbial viability high. In addition, the life span of the ultrasonic transducers is increased and temperature effects due to the transducer usage are minimised [94]. Thus, examination of the effects and the rates of ultrasound intensification utilised appear to be important for individual process strategies.

Reactor design for ultrasonic intensification

The effect of intensification on a bioprocess depends on the choice of ultrasonic parameters utilised. These include the ultrasonic mode, which can be either continuous or pulse, the frequency, the intensity, the processing temperature, the nature of the solvent utilised, the aeration and the design of reactor which determines the level and distribution of energy within the system [19]. The main parameters in ultrasonic reactor design are the type of reactor, its geometry; the design of the transducer set up and the volumetric scale of the feed stock [19]. In addition, the location of the ultrasonic probe in the reacting vessel has been found to influence the distribution of cavitational behaviour within the system [95]. The probe location is usually dependent on the reactor size and the working volume of the liquid reactants [96]. The vertical location of probes in the reactor has been shown to result in poor distribution of cavitation microbubbles with only 10 % distribution reported in a 2.5 L reactor [97]. In contrast, the horizontal location of a probe in a reactor with 82 % immersion length was shown to result in uniform cavitational behaviour over the entire reactor [98]. Another important factor is the shape and diameter of the probe tip. Ultrasonic probes with large diameter tips provide low ultrasonic energy density owing to a wide emitter surface [99]. Usage of larger tips allows higher reaction rates to be achieved. In a study utilising B. amyloliquefaciens producing α-amylase, rapid deactivation of α-amylase was observed to occur, due to the reaction between hydroxyl radicals and the enzyme but this was shown not to take place inside the bubbles. High density cavitational bubbles from a 1 mm sonotrode tip resulted in serious acoustic attenuation and high energy in each cavitational bubble being released when the bubbles collapsed, even though the delivered energy density was low [99]. The radiating face of the ultrasonic probe was shown to greatly affect the efficiency of enzyme inactivation in this case. It is of interest that using large sonotrodes may reduce the intensity of bubble collapse and hence are thus not recommended for intensification [99].

The amount of dissolved gas present in media undergoing sonication is a parameter which also influences the acoustic cavitation. Lowering the quantity of dissolved gas reduces the number of nuclei and thereby increases the cavitation threshold and results in a change in the type of cavitation, producing more vaporous cavities [95]. Such microbubbles collapse less violently and hence the rate of sonochemical reaction will be decreased. The selection of the correct ultrasonic frequency for the applications envisaged is also important. Higher frequency or combinations of lower frequencies is recommended for applications controlled by chemical effects whereas lower frequency is recommended for applications with physical effects [73]. The use of multiple transducers can lead to the formation of uniform cavitational behaviour and also minimise the required energy consumption in applied systems. This occurs as it would not be possible to dissipate the same power through a single probe in a large scale bioprocess operation [95]. In addition, the selection of liquids with low vapour pressure, low viscosity and high surface tension was found to enhance cavitational behaviour [95]. Therefore, the rational design of scaled-up ultrasonic reactors requires the quantitative prediction of acoustic streaming, power dissipation, mass transfer and cavitational activity in the reactor by theoretical simulations [19]. An effective scale-up of the ultrasound reactor can be achieved if the energy dissipation pattern in the reactor is known. One of the major disadvantages of ultrasound baths is the directional sensitivity of the ultrasound waves in the bath, which creates a non-homogeneous energy dissipation pattern. This can be overcome by using tubular reactors with ultrasound intensity concentrated at the core of the reactor, but any industrial scale operation has not yet been reported [100]. In an ultrasonic reactor, the input electrical energy undergoes many transformations while being converted into cavitation energy, which is dissipated in the medium to carry out the physical and chemical change. The cavitation intensity varies with the gas content of the medium [101]. The principles described above have influenced the design of several reactor types for ultrasonication-based processes. Coincidence of the compression half cycles of the two acoustic waves while using dual frequency sonochemical reactor has a favourable effect on the production of radicals, as it intensifies the transient collapse of the cavitation bubbles [102]. The dual frequency ultrasonic processor will not be able to contribute sonochemical effects, which is usually attributed to radical formation during transient cavitation. However, the moderate cavitation intensities in dual frequency reactors would favour the physical processes such as extraction and leaching [103]. Cintas et al. [104] used a reverberative flow reactor, in which two irradiation arrays made up of multiple ultrasonic probes were mounted on the upper and lower portion of a metal cuboid reactor, while liquid reactant was allowed to flow through the cuboid space [104, 105].The reflection of ultrasonic waves at the inner wall of the reactor chamber and its reverberation in the chamber caused the acoustic intensity to be multiplied, thereby giving a more non-uniform cavitation field than single probe systems. Similar technology has also been adopted in sonochemical polygonal reactors [106].

A liquid whistle ultrasonic reactor (frequency 5–30 kHz and intensity, 1.5–2.5 W cm−2) which causes hydrodynamic cavitation has been widely used for industrial wastewater treatment [107], oil emulsification [108], liquid–solid mixture homogenization [19] and fat hydrolysis [109]. This type of reactor can be operated at low cost and is suitable for continuous flow reactions, with scale-up also possible [19, 104]. With effective reactor design as describe, it is possible to maximise the beneficial effects of ultrasonic intensification in a specific process while minimising cost.

Kinetic analyses of ultrasonic intensification processes

The modelling of sonoreactors is challenging due to the experimental issues relating to the effects of ultrasound [110]. The kinetic parameters of ultrasonic intensification on chemical degradation, extraction and fermentation such as ultrasonic frequency, ultrasonic intensity, ultrasonic dose and energy output so far reported are given in Table 5. Ultrasonic Intensity (UI) or acoustic energy density (AED) (sound energy per unit volume) can be evaluated calorimetrically via equation 1 as reported by Mason [111] (Table 5). The influence of the probe surface on ultrasonic intensity can also be determined by the equations 2 and 3 (Table 5). Tiehm et al. [116] hypothesised that the ultrasonic intensity related to the power supplied on the transducer area whereas the ultrasonic density related to the sample volume while the ultrasonic dose related to the energy supplied per sample volume. Though the molecular mechanism of sonochemical degradation remains subtle, there is general agreement that bond cleavage arises from the large shear gradient generated throughout the collapse of the cavitation bubbles. During sonochemical processing, once one goes beyond threshold ultrasound intensity, microbubbles are created which increase in size by absorption of acoustic energy until a critical diameter of 250 µm is reached. Such bubbles become unstable and collapse violently within approximately 20 µs. Adiabatic compression raises the internal pressure to about 500 atm while the cavity temperature may reach 5800 K [117]. If a molecular coil was to be placed beneath the imploding cavity it would be dragged along by the microstream towards the interior of the bubble while being strained by the high fluid velocity gradient [114]. However, the hydraulics are significantly different as the interior of the cavity is filled with a compressible gas with a constantly moving boundary and the applied acoustic pressure is not constant but is a sinusoidal function of time [118]. Nguyen et al. [114] have derived an expression to describe the bubble wall motion during implosive collapse as represented by the strain rate distribution ℇrr using equation 4 (Table 5). Experiments have revealed that the instantaneous radius of the imploding cavity (R) reaches a minimum of the order of 0.5 µm during the final collapse [114]. The strain rate distribution could not be altered other than by changing the ultrasonic conditions. The mechanism of ultrasound assisted acid catalysed transesterification was analysed by the quantitative estimation of physical and chemical effects through simulations of cavitation bubble dynamics. The numerical solution of the bubble dynamics model (see Table 5) can be used to estimate the composition of the bubble contents at the collapse [31]. Ultrasonic intensification can improve the biokinetic parameters such as microbial growth rate [77] during fermentation and also enhance the reaction rate of transesterification during biodiesel production. An investigation of the kinetics of ultrasonic assisted transesterfication of canola waste cooking oil showed that ultrasonication can complete the transesterification within short periods of reaction time and the activation energy was found to be 19, 645 J mol−1 K−1 [119].

Table 5 Kinetic expressions derived for ultrasonic parameters
Full size table

Conclusion

Although research carried out to date has indicated the utility of ultrasonication for enhancement in numerous applications, there remain many factors that require further analysis for a complete optimisation of ultrasonic-based processes particularly with a focus on improving biofuel yields from either extractive or fermentation processes.

Ultrasonic intensification can offer several advantages during bioprocessing; these include low operating cost [120] compared to other enhancing treatment options, simplicity of operation and modest power requirements. In addition, ultrasonic intensification does not require sophisticated equipment reformatting or intensive technical training for utilisation. Ultrasound would likely improve the productivity of many bioprocesses involving live cells via the enhancement of substrate uptake, enhanced production or growth by increasing cell porosity, and potentially enhanced release of cell components which could have important consequences for volatile components such as bioethanol.

Abbreviations

AED:

acoustic energy density

EPA:

eicosapentaenoic acid

UI:

ultrasonic Intensity

References

  1. Sahoo D, Elangbam G, Devi SS (2012) Using algae for carbon dioxide capture and bio- fuel production to combat climate change. Phykos 42:32–38

    Google Scholar 

  2. Chisti Y (2007) Biodiesel from microalgae. Biotech Adv. 25:294–306

    Article  CAS  Google Scholar 

  3. Chisti Y (2008) Response to Reijnders: do biofuel from microalgae beats biofuel from terrestrial plants? Trends Biotechnol 26:351–352

    Article  CAS  Google Scholar 

  4. Norskera NH, Barbosa MJ, Vermue MH, Wijffels RH (2011) Microalgal production—a close look at the economics. Biotech Adv 29:24–27

    Article  CAS  Google Scholar 

  5. Sun A, Davis R, Starbuck M, Ben-Amotz A, Pate R, Pienkos PT (2011) Comparative cost analysis of algal oil production for biofuels. Energy 36:5169–5179

    Article  Google Scholar 

  6. Dexter J, Fu P (2009) Metabolic engineering of cyanobacteria for ethanol production. Energy Environ Sci 2:857–864

    Article  CAS  Google Scholar 

  7. Dexter J, Armshaw P, Sheahan C, Pembroke JT. The State of Autotrophic Ethanol Production in Cyanobacteria. J Appl Microbiol. 2015(In Press)

  8. Gonzalez-Fernandez C, Timmers RT, Ruiz B, Molinuevo-Salces B. Ultrasound-enhanced biogas production from different substrates. In: Fang et al. (eds) Production of Biofuels and Chemicals with Ultrasound. Biofuels and Biorefineries. Springer; 2015; p 209–42

  9. Hansel A, Lindblad P (1998) Towards optimization of cyanobacteria as biotechnologically relevant producers of molecular hydrogen. Appl Microbiol Biotechnol 50:153–160

    Article  CAS  Google Scholar 

  10. Abdel-Basset R, Bader KP (1998) Physiological analyses of the hydrogen gas exchange in cyanobacteria. J Photochem Photobiol B Biol 43:146–151

    Article  CAS  Google Scholar 

  11. Gao Y, Gregor C, Liang Y, Tang D, Tweed C (2012) Algae biodiesel—a feasibility report. Chem Cent J 6(Suppl 1):1–16

    Article  CAS  Google Scholar 

  12. Kaiser BK, Carleton M, Hickman JW, Miller C, Lawson D, Budde M, Warrener P, Paredes A, Mullapudi S, Navarro P, Cross F, Roberts JM (2013) Fatty aldehydes in cyanobacteria are a metabolically flexible precursor for a diversity of biofuel products. PLoS One 8:1–11

    Google Scholar 

  13. Atsumi S, Higashide W, Liao JC (2009) Direct photosynthetic recycling of carbon dioxide to isobutyraldehyde. Nat Biotechnol 27:1177–1180

    Article  CAS  Google Scholar 

  14. Mata TM, Martins AA, Caetano NS (2010) Microalgae for biodiesel production and other applications: a review. Renew Sust Energ Rev 14:217–232

    Article  CAS  Google Scholar 

  15. Harun R, Danquah MK, Forde GM (2010) Microalgal biomass as a fermentation feedstock for bioethanol production. J Chem Tech Biot. 85:199–203

    CAS  Google Scholar 

  16. Ueno Y, Kurano N, Miyachi S (1998) Ethanol production by dark fermentation in the marinegreen alga Chlorococcum littorale. J Ferment Bioeng 86:38–43

    Article  CAS  Google Scholar 

  17. Keris-Sen UD, Sen U, Soydemir G, Gurol MD (2014) An investigation of ultrasound effect on microalgal cell integrity and lipid extraction efficiency. Bioresour Technol 152:407–413

    Article  CAS  Google Scholar 

  18. Lee JY, Yoo C, Jun SY, Ahn CY, Oh HM (2010) Comparison of several methods for effective lipid extraction from microalgae. Bioresour Technol 101(Suppl 1):75–77

    Article  CAS  Google Scholar 

  19. Ensminger D, Bond LJ (2011) Ultrasonics: fundamentals, technologies, and applications, 3rd edn. CRC Press, Boca Raton

    Book  Google Scholar 

  20. Luo J, Fang Z, Smith RL (2013) Ultrasound-enhanced conversion of biomass to biofuels. Prog Energ Combust 41:56–93

    Article  Google Scholar 

  21. Suslick KS, Skrabalak SE (2008) Sonocatalysis. In: Ertl G, Knözinger H, Schüth F, Weitkamp J (eds) Handbook of heterogeneous catalysis. Wiley-VCH Verlag GmbH and Co. KGaA, Weinheim, pp 2007–2017

    Google Scholar 

  22. Ashokkumar M (2011) The characterization of acoustic cavitation bubbles—an overview. Ultrason Sonochem 18:864–872

    Article  CAS  Google Scholar 

  23. Nalajala VS, Moholkar VS (2011) Investigations in the physical mechanism of sonocrystallization. Ultrason Sonochem 18:345–355

    Article  CAS  Google Scholar 

  24. Feng H, Barbosa- Canovas GV, Weiss J (2011) Ultrasound technologies for food and bioprocessing. Springer, New York

    Book  Google Scholar 

  25. Mason TJ (2003) Sonochemistry and sonoprocessing: the link, the trends and (probably) the future. Ultrason Sonochem 10:175–179

    Article  CAS  Google Scholar 

  26. Mason TJ (1992) Practical sonochemistry: user’s guide to applications in chemistry and chemical engineering. Ellis Horwood Ltd., New York

    Google Scholar 

  27. Ranjan A, Patil C, Moholkar VS (2010) Mechanic assessment of microalgal lipid extraction. Ind Eng Chem Res 49:2979–2985

    Article  CAS  Google Scholar 

  28. Balasundaram B, Pandit AB (2001) Significance of location of enzymes on their release during microbial cell disruption. Biotechnol Bioeng 75:607–614

    Article  CAS  Google Scholar 

  29. Kalva A, Sivasankar T, Moholkar VS (2009) Physical mechanism of ultrasound-assisted synthesis of biodiesel. Ind Eng Chem Res 48:534–544

    Article  CAS  Google Scholar 

  30. Khanna S, Jaiswal S, Goyal A, Moholkar VS (2012) Ultrasound enhanced bioconversion of glycerol by Clostridium pasteurianum: a mechanistic investigation. Chem Eng J 200–202:416–425

    Article  CAS  Google Scholar 

  31. Parker PA, Choudhary HA, Mohalkar VS (2012) Mechanistic and kinetic investigations in ultrasound assisted acid catalyzed biodiesel synthesis. Chem Eng J 187:248–260

    Article  CAS  Google Scholar 

  32. Choudhury HA, Malani RS, Moholkar VS (2013) Acid catalyzed biodiesel synthesis from Jatropha oil: mechanistic aspects of ultrasonic intensification. Chem Eng J 231:262–272

    Article  CAS  Google Scholar 

  33. Choudhury HA, Chakma S, Moholkar VS (2014) Mechanistic insight into sonochemical biodiesel synthesis using heterogeneous base catalyst. Ultrason Sonochem 21:169–181

    Article  CAS  Google Scholar 

  34. Choudhury HA, Goswami PP, Malani RS, Moholkar VS (2014) Ultrasonic biodiesel synthesis from crude Jatropha curcas oil with heterogeneous base catalyst: mechanistic insight and statistical optimization. Ultrason Sonochem 21:1050–1064

    Article  CAS  Google Scholar 

  35. Choudhury HA, Srivastava P, Moholkar VS (2014) Single-step ultrasonic synthesis of biodiesel from crude Jatropha curcas oil. AIChE J 60:1572–1581

    Article  CAS  Google Scholar 

  36. Khanna S, Goyal A, Moholkar VS (2013) Mechanistic investigation of ultrasonic enhancement of glycerol bioconversion by immobilized Clostridium pasteurianum on silica support. Biotechnol Bioeng 110:1637–1645

    Article  CAS  Google Scholar 

  37. Singh S, Agarwal M, Sarma S, Goyal A, Moholkar VS (2015) Mechanistic insight into ultrasound induced enhancement of simultaneous saccharification and fermentation of Parthenium hysterophorus for ethanol production. Ultrason Sonochem 26:249–256

    Article  CAS  Google Scholar 

  38. Bharadwaja STP, Singh S, Moholkar VS (2015) Design and optimization of a sono-hybrid process for bioethanol production from Parthenium hysterophorus. J Taiwan Inst Chem Eng. 51:71–78

    Article  CAS  Google Scholar 

  39. Suresh K, Ranjan A, Singh S, Moholkar VS (2014) Mechanistic investigations in sono-hybrid techniques for rice straw pretreatment. Ultrason Sonochem 21:200–207

    Article  CAS  Google Scholar 

  40. Singh S, Bharadwaja STP, Yadav PK, Moholkar VS, Goyal A (2014) Mechanistic investigation in ultrasound-assisted (Alkaline) delignification of Parthenium hysterophorus biomass. Ind Eng Chem Res 53:14241–14252

    Article  CAS  Google Scholar 

  41. Singh S, Sarma S, Agarwal M, Goyal A, Moholkar VS (2015) Ultrasound enhanced ethanol production from Parthenium hysterophorus: a mechanistic investigation. Bioresour Technol 188:287–294

    Article  CAS  Google Scholar 

  42. Gan CY, Latiff AA (2011) Optimisation of the solvent extraction of bioactive compounds from Parkiaspeciosa pod using response surface methodology. Food Chem 124:1277–1283

    Article  CAS  Google Scholar 

  43. Shao Y, Wu Q, Duan J, Yue W, Gu W, Wang X (2014) Optimisation of the solvent extraction of bioactive compounds from LophatherumgracileBrongn. using response surface methodology and HPLC-PAD coupled with pre-column antioxidant assay. Anal. Methods 6:170–177

    CAS  Google Scholar 

  44. Reddy Prasad ADM, Khan (2012) Comparison of extraction techniques on extraction of Gallic acid from stem bark of Jatrophacurcas. J Applied Sci 12:1106–1111

    Article  CAS  Google Scholar 

  45. Liu Z, Ma C, Yang L, Zu Y (2013) Process optimization of ultrasonic-assisted extraction of arabinogalactan from dihydroquercetin extracted residues by response surface methodology and evaluation of Its antioxidant activity. J Chem 2013:1–9

    Google Scholar 

  46. Zhong K, Wang Q (2010) Optimization of ultrasonic extraction of polysaccharides from dried longan pulp using response surface methodology. Carbohyd Polym 80:19–25

    Article  CAS  Google Scholar 

  47. Feng S, Luo B, Tao B, Chen C (2014) Ultrasonic-assisted extraction and purification of phenolic compounds from sugarcane (Saccharumofficinarum L.) rinds. Food Sci Technol 60:970–976

    Google Scholar 

  48. Yong-guang BI, Ding-long Y, Xiao-jun H, Yu-min LI, Min-xia H (2012) Study on ultrasonic-assisted extraction of polysaccharide of Atractylis MacroceohalaKoidz of experiment. Energy Procedia 17:1778–1785

    Article  CAS  Google Scholar 

  49. Ying Z, Han X, Li J (2011) Ultrasound-assisted extraction of polysaccharides from mulberry leaves. Food Chem 127:1273–1279

    Article  CAS  Google Scholar 

  50. Vinatoru M (2001) An overview of the ultrasonically assisted extraction of bioactive principles from herbs. Ultrason Sonochem 8:303–313

    Article  CAS  Google Scholar 

  51. Pingret D, Tixier-Fabiano AS, Chemat F (2013) Ultrasound-assisted extraction. In: Rostagno MA, Prado JM (eds) Natural product extraction: principles and applications. RSC Publishing, Londres, pp 89–112

    Chapter  Google Scholar 

  52. Rocco G, Toledo C, Ahumada I, Sepúlveda B, Cañete A, Richter P (2008) Determination of polychlorinated biphenyls in biosolids using continuous ultrasound-assisted pressurized solvent extraction and gas chromatography-mass spectrometry. J Chromatogr A 1193:32–36

    Article  CAS  Google Scholar 

  53. Lam MK, Lee KT (2014) Scale up and commercialization of Algal cultivation and Biofuel production. In: Pandey A, Lee DJ, Chisti Y, Soccol CR (eds) Biofuels from algae. Elsevier, Amsterdam, pp 261–286

    Chapter  Google Scholar 

  54. Lee AK, Lewis DM, Ashman PJ (2012) Disruption of microalgal cells for the extraction of lipids for biofuels: processes and specific energy requirements. Biomass Bioenerg 46:89–101

    Article  CAS  Google Scholar 

  55. Lee AK, Lewis DM, Ashman PJ (2013) Force and energy requirement for microalgal cell disruption: an atomic force microscope evaluation. Bioresour Technol 128:199–206

    Article  CAS  Google Scholar 

  56. Boyd AR, Champagne P, McGinn PJ, MacDougall KM, Melanson JE, Jessop PG (2012) Switchable hydrophilicity solvents for lipid extraction from microalgae for biofuel production. Bioresour Technol 118:628–632

    Article  CAS  Google Scholar 

  57. Solana M, Rizza CS, Bertucco A (2014) Exploiting microalgae as a source of essential fatty acids by supercritical fluid extraction of lipids: Comparison between Scenedesmusobliquus, Chlorella protothecoides and Nannochloropsissalina. J Supercrit fluids. 92:311–318

    Article  CAS  Google Scholar 

  58. Lee JY, Yoo C, Jun SY, Ahn CY, Oh HM (2010) Comparison of several methods for effective lipid extraction from microalgae. Bioresour Technol 101:75–77

    Article  CAS  Google Scholar 

  59. Keris-Sen UD, Sen U, Soydemir G, Gurol MD (2014) An investigation of ultrasound effect on microalgal cell integrity and lipid extraction efficiency. Bioresour Technol 152:407–413

    Article  CAS  Google Scholar 

  60. Neto AMP, Sontana de Souza RF, Leon-Nino AD, Aparecida da Costa JD, Tiburcio RS, Nunes TA, Sellare de Mello TC, Kanemoto FT, Saldanha-Correa FMP, Gianesella SMF (2013) Improvement in microalgae lipid extraction using a sonication-assisted method. Renew Energ 55:525–531

    Article  CAS  Google Scholar 

  61. Suali E, Sarbatly R (2012) Conversion of microalgae to biofuel. Renew Sust Energ Rev 16:4316–4342

    Article  CAS  Google Scholar 

  62. Suganya T, Renganathan S (2012) Optimization and kinetic studies on algal oil extraction from marine macroalgae Ulva lactuca. Bioresour Technol 107:319–326

    Article  CAS  Google Scholar 

  63. Kaiwan-arporn P, Hai PD, Thu NT, Annachhatre AP (2012) Cultivation of cyanobacteria for extraction of lipids. Biomass Bioenerg 44:142–149

    Article  CAS  Google Scholar 

  64. Sostaric M, Klinar D, Bricelj M, Golob J, Berovic M, Likozar B (2012) Growth, lipid extraction and thermal degradation of the microalgae Chlorella vulgaris. New Biotechnol 29:325–331

    Article  CAS  Google Scholar 

  65. Natarajan R, RusssellAng WM, Chen X, Voigtmann M, Lau R (2014) Lipid releasing characteristics of microalgae species through continuous ultrasonication. Bioresour Technol 158:7–11

    Article  CAS  Google Scholar 

  66. Allard B, Rager MN, Templier J (2002) Occurrence of high molecular weight lipids (C80+) in the trilaminar outer cell walls of some freshwater microalgae. A reappraisal of algaenan structure. Org Geochem 33:789–801

    Article  CAS  Google Scholar 

  67. Geciova J, Bury D, Jelen P (2002) Methods for disruption of microbial cells for potential use in the dairy industry-a review. Int Dairy J 12:541–553

    Article  CAS  Google Scholar 

  68. Park JY, Lee K, Choi SA, Jeong MJ, Kim B, Lee JS, Oh YK (2015) Sonication assisted homogenization system for improved lipid extraction from Chlorella vulgaris. Renew Energ 79:3–8

    Article  CAS  Google Scholar 

  69. Ji J, Wang J, Li Y, Yu Y, Xu Z (2006) Preparation of biodiesel with the help of ultrasonic and hydrodynamic cavitation. Ultrasonics 44:411–414

    Article  Google Scholar 

  70. Sharma YC, Singh B, Upadhyay SN (2008) Advancements in development and characterization of biodiesel: a review. Fuel 87:2355–2373

    Article  CAS  Google Scholar 

  71. Badday AS, Abdullah AZ, Lee KT, Khayoon MS (2012) Intensification of biodiesel production via ultrasonic—assisted process: a critical review on fundamentals and recent development. Renew Sust Energ Rev 16:4574–4587

    Article  Google Scholar 

  72. Zhang X, Yan S, Tyagi RD, Surampalli RY, Valero JR (2014) Ultrasonication aided in situ transesterification of microbial lipids to biodiesel. Bioresour Technol 169:175–180

    Article  CAS  Google Scholar 

  73. Sinisterra JV (1992) Application of ultrasound to biotechnology: an overview. J. Ultrasonics 30:180–185

    Article  CAS  Google Scholar 

  74. Jeon BH, Choi JA, Kim HC, Hwang JH, Abou-Shanab RA, Dempsey BA, Regan JM, Kim JR (2013) Ultrasonic disintegration of microalgal biomass and consequent improvement of bioaccessibility/bioavailability in microbial fermentation. Biotechnol Biofuels 6:1–9

    Article  CAS  Google Scholar 

  75. Chaunyan D, Bochu W, Haun Z, Conglin H, Chuanren D, Wangqian L, Toyama Y, Sakanishi A (2004) Effect of low frequency ultrasonic stimulation on the secretion of riboflavin produced by Ecemotheciumashbyii. Colloid Surf B 34:7–11

    Article  CAS  Google Scholar 

  76. Rapoport N, Smirnov AI, Timoshin A, Pratt AM, Pitt WG (1997) Factors affecting the permeability of Pseudomonas aeruginosa cell walls toward lipophilic compounds: effects of ultrasound and cell age. Arch BiochemBiophys 344:114–124

    Article  CAS  Google Scholar 

  77. Pitt WG, Ross SA (2003) Ultrasound increases the rate of bacterial cell growth. Biotechnol Prog 19:1038–1044

    Article  CAS  Google Scholar 

  78. Choi JA, Hwang JH, Dempsey BA, Shanab A, Min B, Song H, Lee DS, Kim JR, Cho Y, Hong S, Jeon BH (2011) Enhancement of fermentative bioenergy (ethanol/hydrogen) production using ultrasonication of Scenedesmus obliquus YSW15 cultivated in swine wastewater effluent. Energ Environ Sci 4:3513–3520

    Article  CAS  Google Scholar 

  79. Naveena B, Sakthiselvan P, Elaiyaraju P (2012) Partha N Ultrasound induced production of thrombinase by marine actinomycetes: kinetic and optimization studies. Biochem Eng J 61:34–42

    Article  CAS  Google Scholar 

  80. Chisti Y (1999) Mass transfer. Encyclopedia of bioprocess technology. In: Flickinger MC, Drew SW (eds) Fermentation, biocatalysis, and bioseparation. John Wiley, New York, pp 1607–1640

    Google Scholar 

  81. Nyborg WL (1982) Ultrasonic microstreaming and related phenomena. Br J Cancer 45:156–160

    Google Scholar 

  82. Kilby NJ, Hunter CS (1990) Repeated harvest of vacuole located secondary products from in vitro grown plant cells using 1.02 MHz ultrasound. Appl Microbiol Biotechnol. 33:448–451

    CAS  Google Scholar 

  83. Chisti Y (2003) Sonobioreactors: using ultrasound for enhanced microbial productivity. Trends Biotechnol 21:89–93

    Article  CAS  Google Scholar 

  84. Wood BE, Aldrich HC, Ingram LO (1997) Ultrasound stimulates ethanol production during the simultaneous saccharification and fermentation of mixed waste office paper. Biotechnol Prog 13:232–237

    Article  CAS  Google Scholar 

  85. Boateng CO, Lee KT (2014) Ultrasonic-assisted simultaneous saccharification and fermentation of pretreated oil palm fronds for sustainable bioethanol production. Fuel 119:285–291

    Article  CAS  Google Scholar 

  86. Nikolic S, Mojovic L, Rakin M, Pejin D, Pejin J (2010) Ultrasound-assisted production of bioethanol by simultaneous saccharification and fermentation of corn meal. Food Chem 122:216–222

    Article  CAS  Google Scholar 

  87. Yun YM, Jung KW, Kim DH, Oh YK, Cho SK, Shin HS (2013) Optimization of dark fermentative H2 production from microalgal biomass by combined (acid + ultrasonic) pretreatment. Bioresour Technol 141:220–226

    Article  CAS  Google Scholar 

  88. Nguyen TAD, Kim KR, Nguyen MT, Kim MS, Kim D, Sim SJ (2010) Enhancement of fermentative hydrogen production from green algal biomass of Thermotoga neapolitana by various pretreatment methods. Int J Hydrogen Energ. 35:13035–13040

    Article  CAS  Google Scholar 

  89. Schlafer O, Sievers M, Klotzbucher H, Onyeche TI (2000) Improvement of biological activity by low energy ultrasound assisted bioreactors. Ultrasonics 38:711–716

    Article  CAS  Google Scholar 

  90. Avhad DN, Rathod VK (2014) Ultrasound stimulated production of a novel fibrinolytic enzyme. Ultrason Sonochem 21:628–633

    Article  CAS  Google Scholar 

  91. Klomklieng P, Prateepasen A (2012) Molasses fermentation to ethanol by Saccharomyces cerevisiaeM30 using low ultrasonic frequency stimulation. KKU Res J. 17:950–957

    Google Scholar 

  92. Liu R, Li W, Sun LY, Liu CZ (2012) Improving root growth and cichoric acid derivatives production in hairy root culture of Echinacea purpurea by ultrasound treatment. Biochem Eng J. 60:62–66

    Article  CAS  Google Scholar 

  93. Rahman MS (1999) Light and sound in food preservation. In: Rahman MS (ed) Handbook of food preservation. New York, Marcel Dekker, pp 673–686

    Google Scholar 

  94. Raskar HD, Avhad DN, Rathod VK (2014) Ultrasound assisted production of daunorubicin: process intensification approach. Chem Eng Process 77:7–12

    Article  CAS  Google Scholar 

  95. Gogate PR, Sutkar VS, Pandit AB (2011) Sonochemical reactors: important design and scale up considerations with a special emphasis on heterogeneous systems. Chem Eng J 166:1066–1082

    Article  CAS  Google Scholar 

  96. Asakura Y, Nishida T, Matsuoka T, Koda S (2008) Effects of ultrasonic frequency and liquid height on sonochemical efficiency of large-scale sonochemical reactors. Ultrason Sonochem 15:244–250

    Article  CAS  Google Scholar 

  97. Sutkar VS, Gogate PR, Csoka L (2010) Theoretical prediction of cavitational activity distribution in sonochemical reactors. Chem Eng J 158:290–295

    Article  CAS  Google Scholar 

  98. Kumar A, Gogate PR, Pandit AB (2007) Mapping the efficacy of new designs for large scale sonochemical reactors. Ultrason Sonochem 14:538–544

    Article  CAS  Google Scholar 

  99. Kadkhodaee R (2008) Povey MJW. Ultrasonic inactivation of Bacillus a-amylase. I. effect of gas content and emitting face of probe. Ultrason Sonochem 15:133–142

    Article  CAS  Google Scholar 

  100. Moholkar VS, Sable SP, Pandit AB (2000) Mapping the cavitation intensity in an ultrasonic bath using the acoustic emission. AIChE J 46:684–694

    Article  CAS  Google Scholar 

  101. Moholkar VS, Warmoeskerken MMCG (2003) Integrated approach to optimization of an ultrasonic processor. AIChE J 49:2918–2932

    Article  CAS  Google Scholar 

  102. Moholkar VS (2009) Mechanistic optimization of a dual frequency sonochemical reactor. Chem Eng Sci 64:5255–5267

    Article  CAS  Google Scholar 

  103. Khanna S, Chakma S, Moholkar VS (2013) Phase diagrams for dual frequency sonic processors using organic liquid medium. Chem Eng Sci. 100:137–144

    Article  CAS  Google Scholar 

  104. Cintas P, Mantegna S, Gaudino EC, Cravotto G (2010) A new pilot flow reactor for high-intensity ultrasound irradiation. Application to the synthesis of biodiesel. Ultrason Sonochem 17:985–989

    Article  CAS  Google Scholar 

  105. Seymour JD, Wallace HC, Gupta RB (1997) Sonochemical reactions at 640 kHz using an efficient reactor. Oxidation of potassium iodide. Ultrason Sonochem 4:289–293

    Article  CAS  Google Scholar 

  106. Easson MW, Condon B, Dien BS, Iten L, Slopek R, Yoshioka-Tarver M, Lambert A, Smith J (2011) The application of ultrasound in the enzymatic hydrolysis of switchgrass. Appl Biochem Biotechnol 165:1322–1331

    Article  CAS  Google Scholar 

  107. Chakinala AG, Gogate PR, Burgess AE, Bremner DH (2008) Treatment of industrial wastewater effluents using hydrodynamic cavitation and the advanced Fenton process. Ultrason Sonochem 15:49–54

    Article  CAS  Google Scholar 

  108. Shi GB, Shi Y, inventor; Shanghai Shangao Energy Saving Sci and Technology Co Ltd, assignee. Fluidic power ultrasonic emulsifying device useful for heavy oil comprises oil inlet, oil filter, high pressure pump, water tank and filter, oil and water heating device, pre-mixer, and first and second reed whistle ultrasonic emulsifier. Chinese patent: CN201197931-Y; 25 Feb 2009 [in Chinese]

  109. Davidson RS, Safdar A, Spencer JD, Robinson B (1987) Applications of ultrasound to organic chemistry. Ultrasonics 25:35–39

    Article  CAS  Google Scholar 

  110. Grcˇic I, Šipic A, Koprivanac N, Vrsaljko D (2012) Global parameter of ultrasound exploitation (GPUE) in the reactors for wastewater treatment by sono-Fenton oxidation. Ultrason Sonochem 19:270–279

    Article  CAS  Google Scholar 

  111. Mason TJ (1990) Sonochemistry: The uses of ultrasound in chemistry. Royal Society of Chemistry, Cambridge

    Google Scholar 

  112. Neis U, Nickel K, Tiehm A (2000) Enhancement of anaerobic sludge digestion by ultrasonic disintegration. Water Sci Technol 42:73–80

    CAS  Google Scholar 

  113. Choonia HS, Lele SS (2011) β—Galactosidase release kinetics during ultrasonic disruption from Lactobacillus acidophilus isolated from fermented Eleusine coracana. Food Bioprod Process 89:288–293

    Article  CAS  Google Scholar 

  114. Nguyen TQ, Kausch HH (1992) Mechanochemical degradation in transient elongational flow. Adv Polym Sci 100:73–182

    Article  Google Scholar 

  115. Feng X, Lei H, Deng J, Yu Q, Li H (2009) Physical and chemical characteristics of waste activated sludge treated ultrasonically. Chem Eng Process Process Intensif 48:187–194

    Article  CAS  Google Scholar 

  116. Tiehm A, Nickel K, Zellhorn M, Neis U (2001) Ultrasonic waste activated sludge disintegration for improving anaerobic stabilization. Wat Res 35:2003–2009

    Article  CAS  Google Scholar 

  117. Nguyen TQ, Liang QZ, Kausch HH (1997) Kinetics of ultrasonic and transient elongational flow degradation: a comparative study. Polymer 38:3783–3793

    Article  CAS  Google Scholar 

  118. Mason TJ, Lorimer JP (1989) Sonochemistry: theory, applications and uses of ultrasound in chemistry. Ellis Horwood, New York

    Google Scholar 

  119. Grant GE, Gude VG (2014) Kinetics of ultrasonic transesterification of waste cooking oil. Environ Prog Sustain Energy 33:1051–1058

    Article  CAS  Google Scholar 

  120. Thanh LT, Okitsu K, Sadanaga Y, Takenaka N, Maeda Y, Bandow H (2010) A two step continuous ultrasound assisted production of biodiesel fuel from waste cooking oils: a practical and economical approach to produce high quality biodiesel fuel. Bioresour Technol 101:5394–5401

    Article  CAS  Google Scholar 

Download references

Authors’ contributions

This review was conceived, researched and written by Balakrishnan Naveena. Patricia Armshaw participated in the generation of illustrations, discussing the literature and drafting of the manuscript. Joseph Tony Pembroke participated in devising the study, discussions and in the critical review of manuscript draft and revision of manuscript. All authors read and approved the final manuscript.

Acknowledgements

This research was conducted as part through Government of Ireland postdoctoral fellowship (Project Id: GOIPD/2014/303) and funded by Irish Research Council, Ireland.

Compliance with ethical guidelines

Competing interests The authors declare that they have no competing interests.

Author information

Authors and Affiliations

  1. Molecular Biochemistry Laboratory, Materials and Surface Science Institute, Department of Chemical and Environmental Sciences, University of Limerick, Limerick, Ireland

    Balakrishnan Naveena, Patricia Armshaw & J. Tony Pembroke

Authors
  1. Balakrishnan Naveena
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Patricia Armshaw
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. J. Tony Pembroke
    View author publications

    You can also search for this author in PubMed Google Scholar

Corresponding author

Correspondence to Balakrishnan Naveena.

Rights and permissions

Open Access This 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. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Naveena, B., Armshaw, P. & Tony Pembroke, J. Ultrasonic intensification as a tool for enhanced microbial biofuel yields. Biotechnol Biofuels 8, 140 (2015). https://doi.org/10.1186/s13068-015-0321-0

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s13068-015-0321-0

Keywords

  • Algal and cyanobacterial biofuels
  • Ultrasonic intensification
  • Yield enhancement
  • Ultrasonic reactors

Biotechnology for Biofuels and Bioproducts

ISSN: 2731-3654

Contact us