Introduction

The idea of sustainable and innovative bio-economical use of science is the basis for scientific advancements. With more than 4000 different enzymes reported, an average of 200 enzymes has the potential for commercialization, although only 10% can be industrially produced. There is huge potential in the enzyme market, which was reported to be around 6.3 billion dollars in 2017 and has a projection of a compound annual growth rate (CAGR) of 6.8% until 2024. Over the next five years, the food enzyme market is expected to grow by 7.5%, the highest rate of any market projected in the industry (Food enzyme trend gminsight). The thrust to uplift the production of renewable resources is greatly impregnated with the requirement of low-cost yet highly efficient systems (Joshi et al. 2018; Raveendran et al. 2018). White biotechnology is dedicated to harnessing biocatalysts i.e., enzymes and microorganisms at an industrial scale (Meyer et al. 2020, Cairns et al. 2021). The mandate of white biotechnology is to provide pure and replenishable sources as potential alternatives for industrial acceleration resulting in improved, bio-economical, and highly sustainable products (Hyde et al. 2019).

The microbial system is the foundation of biotechnological applications and innovations. The fungal community is a highly exploited eukaryotic system that can be applied directly or by acting as the source to produce industrially important products (Joshi et al. 2018). Filamentous fungi are efficient decomposers that can feed on and break down organic materials and polymeric compounds. Industrial production of commercial citric acid marks the stepping stone of fungal biology as an important commercial industrial product. Enzymes are the backbone of sustainable environments and harnessed industrially for centuries. The fungal system has been a pioneer source of these commercially applicable enzymes. Cellulases, amylases, pectinases, laccases proteases, and lipases are secreted by fungal cells (Hyde et al. 2019; Meyer 2019). These enzymes hydrolyse plant polysaccharides such as cellulose, starch, pectin, proteins, and lipids respectively. Their wide substrate interaction leads to microbial enzymatic intervention in food and feed, pulp and paper, detergent, fuel, pharmaceutical, and chemical sectors (Ahmed et.al. 2021, Kordi et al. 2022).

Pectin alone constitutes approximately 35% of the plant cell wall composition and shows structural complexity and diversity. Pectin polysaccharide gives plant tissues their tensile strength and rigidity. Pectin is frequently employed in the food sector as a gelling agent, thickening, emulsifier, or stabilizer. It can also be used in the pharmaceutical industry as a blood pressure stabilizer, cholesterol controller, and detoxifier (Gawkowska et al. 2018). Natural sources of pectin include fruit waste from pomegranate, banana, lemon, orange, pineapple, wheat bran, malt sprout, and rice bran. Their physiological properties though isolated from different sources remain similar and are beneficial to humankind (de Souza and Kawaguti 2021).

Pectinases are the group of hydrolases, depolymerase, esterases, and lyases enzymes. These act on pectin, protopectin, pectic acid, and galacturonate (Yadav et al. 2009a). Based on the mode of action, specificity of the substrate, and cleavage mechanism there exists a diverse family of pectinolytic microbial enzymes. In this emerging era of biotechnological innovations, fungal pectinase accelerates its way as a promising natural biotechnological innovative agent (Nighojkar et al. 2019; Anand et al. 2020). Microbial pectinases have a broad range of applications and high catalytic effectiveness has significantly raised the global demand. Microbes are natural sources of pectinases that are often employed due to their simplicity of manufacture and distinctive physicochemical features. With a 25% share in the market for food and beverage enzymes, the pectinases family of enzymes is a great part of the biotechnological industry. They are on top of the list of industrial enzymes made for commercial production. Pectinases of acidic nature are preferred for clarification of fruit juices, maceration of vegetables in the manufacturing of pastes and purees, and winemaking. Alkaline pectinases are often used in the retting of natural textile fibres, treatment of pectic-rich wastewater, fermentation of tea, extraction of vegetable oil, and treatment of paper and pulp (Kohli and Gupta 2015; Patidar et al. 2018; Thakur et al. 2021).

The literature available on microbial pectinases has established the importance of the catalyst in industrial sectors. The present review is solely inclined towards fungal pectinolytic interventions in enzyme biotechnology. The status of fungal pectinases and their cost-effective production strategies, the factors affecting production, the large-scale bioreactor-based productions and the purification of the enzyme have been highlighted in this review. Further, emphasis has also been made to include the recent innovations like immobilisation, directed evolution and omics-based approaches targeted in fungal pectinases.

Pectin

Pectin is an abundant natural product predominately observed in dicotyledonous plants. It is secreted by Golgi bodies into the apoplast of cells that are richly methyl-esterified (Sinclair et al. 2018). The committee of the American Society in 1944 accepted definitions of pectic substances, which include pectin acids, pectic acids, and protopectin within the complex class of these macromolecules. The pectinic acids are colloids of galacturonic acids methyl ester and pectic acids without methyl ester (Harholt et al. 2010). Protopectin is considered the parent molecule of a pectic substance and together with pectin and pectic acids was then summarised as “pectin” (Mohen 2008, Anderson 2019).

The solubility behaviour of pectins is categorised as (i) pectins soluble in water or diluted solutions, (ii) pectins soluble in chelators like EDTA, and (iii) protopectin soluble in alkaline or hot solutions based on this observation. The water-soluble and chelator-soluble pectins are derived from the middle lamella of the plant cell wall. These are composed of galacturonic acid residues with a tenth of neutral sugars and barely 2% rhamnose (Voragen et al. 2009; Patidar et al. 2018). The distribution of sugars attached with free carboxyl groups gives the classes their nature of water and chelator solubility. Pectin of alkaline solubility are embedded in part of the cell walls. Alkali-soluble pectins are structured with arabinose and galactose sugars. Typically, softening during ripening or heating is accompanied by a decrease in the proportion of protopectin and an increase in water-soluble pectin (Yapo 2011).

Apart from solubility, pectin has variable percentages of esterification as (i) High-methoxyl pectins, having esterification levels between 40 and 50%, and (ii) Low-methoxyl pectins with esterification levels below 40%. The esterification level can be controlled by acid, alkali, or enzyme treatment of high-methoxyl pectins. This imparts pectin its unique characteristics and helps form gels under specific conditions. The pectin polysaccharide is made up of distinct categories of sugars representing unique structures (Voragen et al. 2009; Wusigale et al. 2020). There are 17 different monosaccharides linked with approximately more than 20 different linkages. These together give pectin a macro-complex structure (Yang and Anderson 2020; Gutierrez-Alvarado et al. 2022). These sugars govern the role of pectin in cell adhesion and separation, expansions and regulation of plant cell walls, and the development of organs and plants.

The complex nature of pectin structure includes components like homogalacturonan (HG), xylogalacturonan (XGA), homogalacturonan, rhamnogalacturonan I (RGI), and rhamnogalacturonan II (RGII) (Yang and Anderson 2020). The key constituent of the spine is α-1,4-linked galacturonic acid (GalA) residues. These residues undergo esterification at six carboxyl carbon and acylation at the third or second oxygen of the chain. This base is alternatively lined by rhamnose sugar and galacturonic acid residues having a structurally similar side chain of arabinose and galactose sugars. Homogalacturonan accounting for 60% of the total pectin structure forms the smooth region of sugar residues. The neutral sugars together are ramified to form a hairy sugar region. Rhamnogalacturonan II (RGII) within HG constitutes twelve different types of sugar resides, including 3-deoxy-lyxo-2-heptulosaric acid (DHA), 3-deoxy-manno-2-octulosonic acid (KDO), apiose and acetic acid. The reproductive tissues, fruits, and seeds store the xylogalacturonan. It forms a single side chain of units of b-D-Xylp-(1 → 3) which is commutated with HG molecules (Zdunek et al. 2021; Shin et al. 2021; Gutierrez-Alvarado et al. 2022).

Pectinases family: an overview

Pectinases act on pectic substances. They possess negative charge, high molecular weight glycosidic bond-linked macromolecules with substrate specificities on pectin (Anderson 2019). Pectinases are classified in respect of the type of modifications of the backbone chain as protopectin, pectic acid, pectin acid, and pectin. Pectinases amalgamate together lyases, hydrolases, and esterases classes of enzymes to act on pectin (Yadav et al. 2009b, Pedrolli et al. 2009). These can work endogenously by cleaving glycosidic bonds to release residues from the inside or in an exogenous manner to cleave residues from the ends. These can be produced through extracellular or intracellular modes. Though intracellular secretion is more costly in comparison to extracellular production. The classification of pectinases or pectinolytic enzymes based on the existence of different pectic substances, reaction mechanisms, and degradation of the hairy and smooth regions has been reported (Kashyap et al. 2001; Jayani et al. 2005; Favela-Torres et al. 2006).

A discrete collection of two hundred and sixty-nine enzymatic families that are similar on grounds of amino acid sequence are called the Carbohydrate-modifying enzymes. This family is broadly distributed under four classes: glycoside hydrolases (GHs), glycosyltransferases (GTs), polysaccharide lyases (PLs), and carbohydrate esterases (CEs). These classes have subgroups of structurally and catalytically related families. This has been listed in the carbohydrate-active enzyme (CAZy) database (www.cazy.org) (Cantarel et al. 2009). Pectinases share a diverse group of enzymes that distinctively occupy their positions in the GH, PL, and CE families (Drula et al. 2022).

Glycoside hydrolases (GH)

Family GH28 is commonly referred to as polygalacturonases, which are glycosidases acting on homogalacturonan and rhamnogalacturonan components of pectin. This includes enzymes with hydrolysis mechanisms. They are capable of hydrolysing glycosidic linkage between carbohydrates -carbohydrates and a non-carbohydrate moiety. GH28 enzymes are also categorized into three distinct categories acting on homogalacturonan, rhamnogalacturonan and xylogalacturonan (Sprockett et al. 2011; Villarreal et al. 2022). It hydrolyses polygalacturonic acid on α-1,4-glycosidic linkages producing d-galacturonate. Fungal polygalacturonase can produce monomeric galacturonic acids on its depolymerization. Mode of action distributes them as Endo-PG (EC 3.2.1.15) which liberates saturated oligogalacturonides and Exo-PG (EC 3.2.1.67) releases saturated galacturonic acid residue. The residue is obtained from the non-reducing end of homogalacturonan by hydrolytic catalysis (Yang et al. 2018; Anand et al. 2020; Christensen 2020). Xylogalacturonans (XG) are enzymes responsible for cleaving glycosidic linkages in the xylose-substituted rhamnogalacturonan chain and the end products are xylose-galacturonate dimers. Rhamnogalacturonan is hydrolytically cleaved by RG galalcturonohydrolase. Its non-reducing end produces monogalacturonate (Villarreal et al. 2022).

Polysaccharide lyases (PL)

These enzymes cleave uronic acid-containing polysaccharide chains. They use the β-elimination mechanism to generate an unsaturated (hexen)uronic acid residue and a new reducing end. PLs, can cleave alginate, heparin, hyaluronan, pectin, xanthan, and several exopolysaccharides (cazy.org/Polysaccharide-Lyases; Yadav et al. 2009c, Chakraborty et al. 2017). PL family 1, 2, and 9 share distributions of lyases degrading pectin. Pectate lyase (PL) results in forming an unsaturated product (α-4,5-d-galacturonate) through a trans-elimination reaction on polygalacturonase acids. Endo-PL (EC 4.2.2.9), acts on a nonreducing end. Pectin lyase (PNL) (EC 4.2.2.10) results in the formation of 4,5- unsaturated oligo-galacturonate. PNL performs a β-elimination mechanism without affecting the ester content of the polymer chain. This ester content is responsible for the specific aroma of fruits. Toxic methanol production is limited by enzymatic degradation. Henceforth, these are preferred in fruit juice clarification industries. Rhamnogalacturonan lyases degrade rhamnogalacturonan I and are distributed in families 4 and 11 (Zheng et al. 2021).

Carbohydrate esterases (CE)

These enzymes catalyse acylation at the oxygen or nitrogen end. The members of this family remove esterified modifications from mono-, oligo- and polysaccharides. The acylation provides easy access to glycoside hydrolase (cazy.org/Carbohydrate-Esterases; Wardman et al. 2022). Pectin methyl esterases are grouped in CE 8 family and act preferentially on a methyl ester group of galacturonate to produce methanol and pectic acid. The action of PMEs forms pectate gel from homogalacturonan. The action of esterases can hinder the action of polygalacturonases (Benen et al. 2002). Rhamnogalacturonan acetyl esterase is responsible for cleaving acetyl groups of the rhamnogalacturonan chain that constitutes the major part of the hairy portion of pectin and belongs to the family CE12. Pectin acetyl esterase belongs to CE13 and hydrolyses the acetyl ester of pectin. They help the formation of pectic acid and acetate and acylation affects the age and differentiation of plant tissues. It even acts as protection from different enzymatic interactions. The esterases assist actively in biomass saccharification and have diverse biological and biotechnological applications (Benen et al. 2002; Bonnin and Pelloux 2020). The diversity of pectinases and their potential for industrial application is depicted in Fig. 1.

Fig. 1
figure 1

Pectinases: classification and role as industrial catalyst

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Fungal pectinases

Microorganisms have been in the environment from the beginning of time on this planet. In the scientific world, the study of the structural, functional, and ecological attributes of microorganisms is significant (Prasad et al. 2021). Microbial enzymes particularly from fungi are preferred over other sources because: (i) Their content is more predictable. (ii) They have a wide range of enzymes; (iii) Bulk production generally resin with low costs and reliable raw materials. (iv) Their productivity rate is high and they contain a greater amount of active ingredients. (v) Fungi can be easily managed to take the desired enzymes, and they can be made in large quantities rapidly and inexpensively through existing fermentation techniques and sophisticated instrumentation. (vi) Enzyme production may be programmable in various environments. and (vii) more potentially hazardous components like phenolic compounds, endogenous enzyme inhibitors, and proteases are found in plant and animal tissues than in microorganisms (Sharma et al. 2013; Singh et al. 2019).

Fungal microbiota cover about 50% of microbial enzyme production. Around 35% of the production shared is held by bacteria and only 15% is produced from higher organisms (Wösten 2019). The fungal system is a popular source of enzymes because they provide a cost-effective technology with reduced resource consumption and minimal emissions, as opposed to animal and plant sources. Fungi and yeast alone are a producer of half of the globally used enzymes (Lübeck and Lübeck 2022). The secretion of pectinases by fungi assists in the breakdown of the middle lamella in plants (Prasad et al. 2021). Soil is a diverse and dynamic environment that is home to a diverse range of microorganisms, especially fungi. The traditional laboratory culture techniques have made the greatest contribution to gaining access to microbial diversity. Despite its familiarity and use, it is one of the world’s least explored environments. Soil microorganisms play a crucial role in plant development and carbon and nutrient cycling. The bulk of soil microorganisms, on the other hand, have yet to be isolated, and their roles are mostly unknown. These microbial communities are exploited as a sustainable source for microbial enzymatic production systems (Baldrian 2019; Selvasekaran and Chidambaram 2020).

Fungi generate a plethora of extracellular enzymes capable of degrading organic materials, one of which is pectinolytic enzymes. Commercial enzymes have been produced using filamentous fungi for more than 50 years (Haile and Ayele 2022). Pectinolytic enzymes are one of the extracellular enzymes that fungi produce that can break down organic molecules. One of the most potent sources of pectinases is filamentous fungi, which can be extensively exploited in the production of SSF at a low cost. Many different fungal species have been reported to produce pectinases. Aspergillus niger is the most typical fungus used in the production of pectinolytic enzymes for industrial use (Gutiérrez-Correa et al. 2012). A. oryzae, A. fumigatus, A. terreus, A. sojoe, A. awamori, and other Aspergillus species are also known to produce pectinase. A. giganteus was the first species whose production of endo-PGL was noted. Additionally, species of Penicillium, Fusarium, Mucor, Neurospora crass, Sclerotinia sclerotium, and others play a part in the manufacture of pectinase (Sharma et al. 2013; Haile and Ayele 2022). Fungal pectinases play a part in the phytopathological process. They interact in plant–microbe symbiosis, and the decomposition of dead plant material, thereby, contributing to the natural carbon cycle. In the context of mining fungal pectinolytic sources, soil samples have been indefinitely explored for the isolation of novel fungal strains as listed in Table 1.

Table 1 Fungal strains isolated from different soil samples with potential for pectinase production using fermentation methods
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Production strategies

Fermentation-based production of microbial pectinases is facilitated by solid-state fermentation and submerged fermentation industrially and at a small scale. The advantages of fermentation-based production of enzymes include low costs, low energy consumption, and low waste-water generation, and it can be exploited to repurpose organic wastes into value-added products. Fermentation-based microbial enzyme mass production uses either solid-state fermentation (SSF) or submerged fermentation (SmF). SmF technology is often used to produce microbial enzymes, especially from bacterial sources and the major advantage is easy to control the process as compared to SSF (Sharma et al. 2013).

Solid State Fermentation uses a solid substrate that acts as a natural habitat for fungi to attach. The fermentation requires lower to no moisture content occurring in the absence or near absence of free water. The sturdy foundation offers support, or occasionally both support and sustenance. The main benefits of SSF include low capital expenditure, reduced levels of catabolite repression and end-product inhibition, low wastewater output, improved productivity, higher enzyme yields, and better product recovery. SSF has been used predominantly as it triggers the production of various enzymes directly from raw materials rich in lignocellulose (Kumar and Verma 2020). SSF is highly favourable for fungal microflora as it is like their natural habitat. Some of the limitations of the SSF include the need for proper aeration and humidity control and a time-consuming scale-up process. Pectinases of fungal origin have been extensively reported by using the solid-state fermentation method (Soccol et al. 2017; Lizardi-Jiménez and Hernández-Martínez 2017). The production of fungal pectinases by SSF requires optimization of several parameters which can directly affect the enzyme production.

Factors influencing the production of pectinases by SSF

The process of fermentation is dependent on biological and physio-chemical parameters that greatly affect the kinetics of the microbial enzymes. To improve the efficiency of the enzymes, these parameters need to be optimised and microbes, the size of the inoculum, and substrates are some of the important biological parameters. Further, incubation temperatures, pH specificities, moisture content, aerations, rotations, and heat transfer affect the performance of the enzymes (Soccol et al. 2017).

Fungal spores can directly be added as inocula and have a very fast production rate. These can grow over a range of temperature conditions between 24 and 30 °C. Thermophilic fungi can also grow optimally in this range. The pH range may change according to the substrate used, however, for the best growth, fungi strains prefer an acidic to a neutral range (Prado Barragán et al. 2016). Optimizations of moisture content have resulted in more stress-resistant pectinase production. The rotation and agitation affect microbial growth and contamination. Production of fungal pectinase is severely constrained by bacterial contamination (Prado Barragán et al. 2016; Chen and Wang 2017).

The mesophilic and thermophilic fungal pectinase production during SSF is also affected by heat transfer during the process. The gases produced by the fungal inoculum and moisture vaporization regulate the heat of the system (Kumar et al. 2021; Chilakamarry et al. 2022).

Substrates used for the production of fungal pectinase

Higher fungi have well-tuned enzymes, spores, and metabolites for development on solid, moist substrates. For instance, fungus spores produced by SSF display greater stability, are more resistant to drying, and have higher germination rates for longer periods (Arun et al. 2020). The substrate acts as structural ed support rich in nitrogen and carbon for the growth of microorganisms. The nutritional composition and quality problems may affect the fermentation batches. This variation could lead to decreased production. The choice of substrate will determine how much heterogeneity is introduced during the process. The most often utilised substrates for SSF include agricultural and food processing wastes such as wheat bran, sawdust, apple pomace, cassava, sugar beetroot pulp, citrus waste, maize cob and banana waste. Innovations in the production of pectinases using different agro-wastes like peels and pulps of citrus, orange, coffee, grapefruit, and banana using both SSF and SmF have been reported recently (Bharathiraja et al. 2017; Chilakamarry et al. 2022).

Fruits and vegetable peels are rapidly utilized nowadays as they are environment-friendly and immensely nutritious for microbes. Peels of citrus fruits, bananas, sweet potatoes, and mango are being vigorously studied. The pomace of apple, kiwi, peach, and grapes are pectin-rich biomass for valorisation via fermentation. Other agro-industrial residues such as oil cakes of pumpkin, sesame, groundnut, and sunflower oil have also been used as substrates (Lopes and Ligabue-Braun 2021). Additionally, pectinolytic enzyme production has been reported by the use of sugarcane bagasse, corn cobs, soybean hulls, sugar beetroot pulp, barley husks and straws as sources of carbon. Tea extract serves as an important source of nitrogen. In addition to these, brewery waste, sewage wastewater, drainage effluents, tobacco stalks, molasses, and vegetable and fruit juices work excellently as liquid substrates for the fermentation of fungal pectinases (Sadh et al. 2018; Cano et al. 2020; Chukwuma et al. 2020).

Bioreactors for the production of fungal pectinases

For large-scale bulk production, bioreactors have been used. These bioreactors or fermenters are designed for processing biological products under a specifically controlled environment. Bioreactors for fungal pectinases have used lignocellulosic wastes, and agricultural wastes as substrates for industry efficient scaled production of enzymes (Cerda et al. 2019). Bioreactors prefer solid state-based fermentation methods for the production of fungal pectinases. In the light of fungal pectinases, Aspergillus niger has been extensively utilized for pectinases production by solid-state fermentation using the packed bed, and bench scale rotating-drum reactors (Finkler et al. 2017; Poletto et al. 2017; Reginatto et al. 2022). A 40 cm high packed bed bioreactor yielded productivity of 1840 U/g pectinases using Aspergillus niger (Pitol et al. 2016). Raimbault columns, packed-bed bioreactors, Erlenmeyer flasks, perforated trays, and other static bioreactors have been used to produce pectinases (Yang and Sha 2019). These bioreactors are chosen because of their usability and simplicity. A. niger on sugarcane bagasse and orange pomace has been utilized as solid-state substrates for production using a tray and rotating drum bioreactors (Mahmoodi et al. 2019). Agitated bioreactors utilise intermittent or continuous mixing to homogenise substrate using solid-state fermentation. It is possible to construct agitated bioreactors with or without a water jacket to regulate temperature (Mitchell and Krieger 2019). This type of reactor may be continuously or intermittently agitated. Shear problems and damage to the fungal mycelium’s structural integrity may occur, depending on the degree of agitation (Shanmugam et al. 2022). Large fermenters are commonly built of stainless steel in the food and beverage industries because of their ability to resist corrosion. A bioreactor’s design incorporates numerous essential engineering elements that are regularly updated and modernised to increase the final product’s productivity and quality (Kaur and Kaur 2019). Basal stirred tank fermenters utilised A. foetidus strain for optimization and evaluation of pH effect on microbial enzymes including pectinases (Li et al. 2018). Innovative forms of bioreactor-based fermentation largely depend on aeration techniques. These reactors have been modernised with the inclusion of steam traps, valves, mechanical foam breakers, pH temperature and pO2 monitors, micro-spargers for self-cleaning, and other sampling ports. Connecting to computers is a crucial advancement for novel bioreactors since it speeds up data processing and calculation and facilitates operational optimisation (Mitchell et al. 2019; John et al. 2020; Leite et al. 2021).

Response surface methodology (RSM) utilisation for bioreactor-based production using shake flasks has been utilised recently to produce pectinases at a concentration of 380 U/ml by A. sojoe (Fratebianchi et al. 2017). Similarly, an indigenous Aspergillus sp. isolated from coffee waste was used in response surface methodology designed on an SSF-based tan ray bioreactor to yield 29.9 IU/g of pectinases (Núñez Pérez et al. 2022).

Purification of fungal pectinases

Enzyme purification can be achieved by using a variety of conventional and modern techniques. The choice of the best treatment stage is a prerequisite for the enzyme purification process to be successful. Depending on the intended usage of the enzyme, the degree of purification may vary. Purification of microbial pectinases has been attained by simple centrifugation, sedimentation, or precipitation (Holm et al. 2018). The removal of inorganic and organic impurities is highly feasible by salting out using ammonium sulphate salts. This method of purification or partial purification has yielded a stable protein with better activity. Solvent precipitation using acetone, ethanol, and methanol, based on the solubility of protein is a cost-effective method for the removal of organic and inorganic impurities. The salt-based precipitation has been preferred as other solvent methods for pectinase. This is generally followed by dialysis to yield salt unbound proteins which are dissolved in buffers for optimal activities. Purification using counter solvents like butanol or octanol or by ultrafiltration facilitates the generation of aqueous pectinase. This eliminates the need for precipitation with dialysis of salt-based methods (Patel et al. 2017; Raina et al. 2022).

Purification of pectin lyases produced from Penicillium oxalicum, P. citrinum, Aspergillus flavus, A. ficuum, A. terricola, Fusarium decemcellulare, and F. lateritum has been performed simply by using ammonium sulphate precipitation and column chromatography method (Yadav and Shastri 2007; Yadav et al. 2008, 2009a, c, 2013, 2014, 2017b). Exo-polygalacturonase from Aspergillus flavus has been purified using solvent-based acetone purification, followed by cellulose column and gel filtration chromatography (Anand et al. 2017a).

Ion exchange, gel filtration, and affinity-based chromatographic methods are used to produce samples with a comparatively greater level of purity. The form, size, charge, hydrophobicity, or binding ability of the stationary phase are criteria used in chromatographic procedures to purify microbial pectinases. The molecular properties and interactions that underlie ion exchange, surface adsorption, partition, and size exclusion are also important considerations (Coskun 2016). Pectinolytic purification has been predominately accomplished by column chromatography (Smith 2005; Ullah 2012; Bassim Atta and Ruiz-Larrea 2022). Ion exchange or gel filtration, which gives rise to purer fractions of pectinases, along with a significant increase in its specific activity has also been reported. Anion exchange column-based purification for polygalacturonase from Calonectria pteridis utilized eucalyptus leaves in submerged fermentation (Ladeira Ázar et al. 2020). An indigenously isolated soil-borne Aspergillus japonicus yielded 2.9-fold purified polygalacturonase using two chromatographic techniques simultaneously (Cavalieri de Alencar Guimarães et al. 2022). A repertoire of purification strategies has been adopted for the purification of fungal pectinases from different fungal strains as shown in Table 2.

Table 2 Purification and biochemical characterization of Fungal Pectinases
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Innovations: diverse approaches

Immobilisation

The pectinolytic industrial intervention is disrupted due to their recovery rates, and low stability. Immobilization of enzymes enhances storage, reduces product contamination, and simplifies the separation of products, which in their free form is challenging. It improvises the catalytic properties of enzymes and enhances their functioning in adverse conditions (Bashir et al. 2020). Thereby, facilitating the recovery and reuse of enzymes in the medium and enhancing the economic feasibility of the enzymes. Suitable immobilization protocols and supportive environments are required for enzyme biocatalysts with high enzymatic activity (Patel et al. 2022). Pectinases have been immobilized using diverse supports by membrane adsorption, covalent binding, and cross-linking mechanisms. A variety of supports, including beads, microspheres, pulp fibre, matrix, resins, capsules, nanoparticles pumice, and magnetic beads have been deployed (Martín et al. 2019; Karataş et al. 2021). The magnetic core of magnetic particles as beads makes it simple, rapid, and effective to separate the enzyme from the reaction mixture using an external magnetic field, making them suitable support for enzyme immobilization. Additionally, the size of the particle can be adjusted to give a large surface area and high enzyme activity (Soozanipour et al. 2019; Trindade Ximenes et al. 2021). Direct crosslinking of different enzyme preparations is the most typical technique for producing cross-linked enzyme aggregates (CLEAs). The advantages of this approach are highly concentrated enzyme activity, greater stability, and the absence of an extra carrier’s associated production costs (Nouri and Khodaiyan 2020).

Adsorption, covalent binding, and entrapment are just a few of the methods utilised to keep enzymes inside the membrane. Enzymes are frequently attached to membranes by chemical bonds and adsorption. Pectinase is frequently bound to membranes using adsorption techniques. Chemical enzyme binders including glutaraldehyde, glycidyl methacrylate, and carbonyl diimidazole are used to adsorb membranes. It has been observed that membrane-bound enzyme exhibits enhanced thermal stability and temperature optima. Among the different methods of immobilising enzymes, covalent immobilisation is frequently preferred. This is so that it won’t allow the enzyme to desorb from the support during the process (Nadar and Rathod 2019).

A scale bioreactor used in stainless steel bases matrix was immobilized to get a titre of 307.5 and 242.6 U/ml of exo and endo PG respectively from Rhizopus oryzae (Zheng et al. 2017). Beads of alginate-montmorillonite were used to immobilize pectinase from A. aculeatus recovering 53% of its initial activity (Mohammadi et al. 2019). Gel-based beads of alginate and agar facilitate the immobilization of pectinase from A. awamori. This retained initial activity even after 8 cycles of reaction (Abdel Wahab et al. 2018). An indigenously isolated pectinolytic yeast strain, Geotrichum candidum was immobilized retaining 70% of its initial activity using corn cob matrix (Ejaz et al. 2018). Similarly, beads of sodium alginate were used in different strains of Geotrichum candidum to immobilize pectinase enhancing its activity from 0.046 to 0.115 IU mL−1 (Ejaz et al. 2020). Pectinases have also been immobilized using magnetic chitosan particles by direct extraction from fruit juices without the intervention of microbes (Dal Magro et al. 2018, 2019; Soozanipour et al. 2019). Efforts on the immobilization of pectinases from fungal strains have been summarized in Table 3.

Table 3 Reports on immobilisation of Fungal Pectinases
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Directed evolution

The state-of-the-art technology of directed evolution for the desired manipulation of enzymes for industrial application has been attempted for pectinases. Mutation using a UV range of 254 nm has been used for the enhancement of polygalacturonases production of Aspergillus and Penicillium species (Heerd et al. 2014; Kamalambigeswari et al. 2018; Nawaz et al. 2019). Mutated strains have also been used to study evolutionary relationships between PEL and PL subclasses of pectinases (Yang et al. 2020). Mutation of gaaX and gaaR allowed A. niger to express pectinases without an inducer (Alazi et al. 2019). The approach of directed evolution combined with computational technologies has been used to access different metabolic pathways of fungal pectinases (Wang et al. 2021). For fungal pectinases, artificial environments can be simulated through strain mutation, recombination, and gene overexpression. With this modification, the pectinolytic mechanism can be accelerated to catalyse chemical reactions in an entirely new environment employing a newer substrate, resulting in increased catalytic activity. Chromosomal mapping was used to analyse S. bayanus var. uvarum strains, and the results revealed three divergent genes, PGU1b, PGU2b, and PGU3b, which are situated on chromosomes X, I, and XIV, respectively. As a result, it was demonstrated that these yeasts’ strong pectinolytic activity might be caused by the existence of many PGU polymeric genes in their genomes (Naumova et al. 2019). Heterologous expression of fungal pectinase targeting expression using microbes with a high capacity for protein production and enzyme secretion has been performed. It is a good alternative to the fermentation technique for the desired production of enzymes by targeting the relevant genes. The expression of pectinolytic genes has been summarized in Table 4.

Table 4 List of pectinase gene studies
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Omics interventions

The omics-driven approach is the current trend in enzyme research which aims to analyse the potential of fungal species in terms of enzyme production by targeting the whole genome or proteome. Over 50% of the currently available eukaryotic genome sequences are from the kingdom of Fungi. Several fungal genome sequences have been targeted to decipher the diversity of pectinases. Recently using a shotgun proteomics approach two pectin lyase and one pectate lyase from Saccharomyces cerevisiae produced using passion fruit flour by solid-state fermentation has been reported (Takeyama et al. 2022). Two-dimensional electrophoresis-based proteomic analysis of Aspergillus niger EIMU2 has been attempted. It revealed that the mutant EIMU2’s multiple enzyme systems used for the degradation of pectin included the main-chain cleaving enzymes polygalacturonase, pectate lyase, and pectin esterase, as well as some accessory enzymes rhamnogalacturonan lyase (Lin et al. 2021). Studying the interaction of wood rotting fungi, pectinases proteomics profiling helped analysed other proteins secreted which might have a significant role in degrading wood (Presley et al. 2020). CRISPR/Cas9 system generated three chimeric GaaR-XlnR induces by D-galacturonic acid from Aspergillus niger. Their proteomics investigation verified that the gaaR mutants carrying the chimeric transcription factor produced several pectinolytic enzymes (Kun et al. 2021). The PL7 and PL8 enzymes required for the breakdown of laminarin, cellulose, lipids, and peptides, were found to be abundantly secreted by Paradendryphiella salina cultured on brown algae using proteomic analysis (Pilgaard et al. 2019). However, a significant issue with the existing fungal pectinases proteomics is to fully understand the expression, operation, and regulation of the entire set of fungus-genome-encoded proteins. Moreover, the sequencing of several fungal proteomes is in progress (Sudhakar et al. 2018).

Meta-omics approach collects total environmental DNA which is targeted for metagenomic studies. A metagenomic system can be any arbitrary environmental sample defining the collection of microbes. Soil, water, air, cow rumen, and composts are such systems, thus, opening doors for unculturable and novel sources for catalytic enzymes. metagenomic approach for pectinase enzyme mining from soil resulted in the isolation of thermostable pectinase (Singh et al. 2012a, b). This approach has been used for identifying novel fungal sources for pectinases (Tanveer et al. 2016; Pilgaard et al. 2019; Ahmad et al. 2021). The metagenomic studies exclusively for fungal pectinases are summarized in Table 5.

Table 5 Metagenomic intervention in search of novel Fungal pectinases
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Industrial applications

Pectin in plant cells is degraded by pectinases. They were first used commercially in the 1930s, and since then, they govern 25% of industrial applications. Wide-ranging industrial uses for pectin-degrading enzymes include degumming and retting of plant fibres, oil extraction, fruit juice clarification, wine production, fermentation of tea and coffee, bioconversion of wastes, and protoplast fusion technology (Singhania et al. 2015). Since 40% of the dry weight of plant cambium cells is made up of pectin, pectinases are essential for digesting natural fibres. With the aid of pectinases, the bast fibres of jute, flax, hemp, ramie, banana, pineapple leaf, and bamboo can be successfully degummed, macerated, and retted because they break down the pectin in the middle lamella and primary cell walls. Their wide applicability in the textile industry makes their study essential. Microbial pectinases-based natural fibre retting and extraction is biodegradable, recyclable, cuts production costs and is energy sustainable (Kumari et al. 2021). The fibres produced are reported with higher strength, shinier, easy to obtain and light weighted. The increasing demands on enzyme applications are growing as replacements for traditional harsh chemical processes. Fungal pectinases are also used for degumming natural fibres, bio scouring, bio bleaching and in wastewater treatment of textile power plants (Sharma et al. 2017).

They are also used to produce effective viral preparation from plant tissues, in the treatment of wastewater and for the isolation of protoplasts. Protoplasts are isolated from the mycelia of Pleurotuseous and Pleurotus flabellatus using enzymes comprising commercial cellulases, crude pectinases, and crude chitinases (Eyini et al. 2006; Ruiz et al. 2017). Pectinases are also applied in animal feeds as it helps in the efficient absorption of nutrients by animals by degrading the fibres that entrap them. These groups of enzymes have been used for biofuel production like bioethanol. The rate of ethanol generation rises when pectinaceous structures in the feedstock are destroyed and hydrolyzed by pectinases. Biomass enzymatic hydrolysis is a cost-effective and efficient treatment method that produces no hazardous waste (Samanta 2019). Sugar becomes more accessible and sensitive to hydrolytic enzymes after being treated with liquid hot water. Alkaline pectinases both from fungal and bacterial sources are also applied in the fermentation of coffee and tea. Degrading pectin, pectinase increases the pace of tea fermentation and reduces the foaming ability of instant tea granules (Tatta et al. 2022).

The fruit and food processing industries have wide applicability of pectinases. Fruits have a complicated pectin structure, making it challenging to extract juice from this very viscous, jellified pulp (Pagnonceli et al. 2019). The pectinase enzyme acts on the pectin of fruit peels and dissolves the glycosidic linkages between the galacturonic acid monomers, reducing the amount of water that may be held by pectin enzymatic treatment is the most frequently used method for juice extraction and clarity (Anand et al. 2017b). The enzymatic hydrolysis of cell walls enhances the extraction yield, soluble dry matter content, galacturonic acid content, and titratable acidity of the products. The amount of waste pomace decreased and the resulting pulp had a lower viscosity. The biomaterial is enzymatically degraded depending on the type of enzyme, incubation period, temperature, concentration, agitation, pH, and the use of various enzyme combinations. The wine industry chooses pectinases as they increase wine quality, and facilitate extraction, filtering, and taste and colour intensification (Gunjal et al. 2020). Pectinases were also used in extracting essential oils from a variety of sources like olives, flaxseed oil, dates, and other fruits and vegetables (Nagpal et al. 2021). These enzymes help to enhance the fatty acids, peroxide value, and colour intensity as compared to chemical treatment. In the paper industry, pectinases along with xylanases are preferred as a bio-bleaching agent. Enzymatic intervention is eco-friendly, less abrasive, and effective in improving paper quality (Nagpal et al. 2020). Biological bleaching with pectinases and xylanases brightens the paper and improves its physical characteristics, as well as lowers the kappa number and permanganate number of the pulp. In comparison to those chemical alternative solutions, the substitution of pectinases contributes to a reduction in chlorine discharge into the environment (Nagpal et al. 2020; Tatta et al. 2022). The diverse industrial application of pectinases has been summarized in Table 6.

Table 6 Potential industrial applications of Fungal pectinases
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The bottom line and future prospects

Pectinases represent an important group of enzymes with immense potential for diverse industrial applications. Substantial efforts have been made to explore the possibility of diverse approaches for enhancing pectinases production, manipulation and elucidating industrial applications, exclusively from fungal sources. The cost-effective production of fungal pectinases using agro-wastes is an eco-friendly approach that has immense potential for converting waste biomass. It also results in the production of different value-added products. This is also added to the saccharification potential of pectinases. Efforts have been made to optimize growth conditions as a precursor to enhanced fungal bioproduct production. Utilising waste valorisation techniques, it is possible to take advantage of the diversity of fungi by using contaminated items as a source of fungi. The fungus system offers many advantages and benefits, but it also poses a hazard due to its pathogenicity and ability to mitigate spoilage and damage. Recombinant and mutagenic approaches can be used to change the pathogenicity of native fungus hosts. According to industrial needs, the fusion of traditional and modern state-of-the-art technology has enormous potential.

Over the years, several fungal genera have been targeted for the production of pectinases and efforts have been made to enhance the catalytic activity, specificity, and applicability for industrial applications. Dual culture inoculums for fermentation-based manufacturing have been employed to increase enzyme productivity. These involve using more than one fungal species for the production of the same biocatalyst. But they strictly demand more comprehension of how various hosts interact with one another. The metagenomics approach has resulted in the deciphering of novel microbes with enhanced pectinase activity, thereby giving the world new industrially potent species. Despite metagenomics inclination in microbial studies, fungal metagenomic library construction and diversity studies are minimal. Though purity of metagenomic DNA from humic acid contamination and the easy extraction of prokaryotic diversity in metagenomics DNA limits the studies of pectinases of fungal metagenomic origin from s potential. The directed evolution approach for altered pectinases activity and specificity has resulted in diverse industrial applications predominately in the textile and food industries. Omics-driven approaches including genomics, proteomics, and metabolomics have been used for understanding the production and expression of pectinase genes. Sequencing of fungal strains, genome-wide mining of pectinases using a bioinformatics approach, and expression of the identified pectinases are intensely investigated areas of research in fungal pectinases. Immobilisation of fungal pectinases using novel approaches for enhancing stability and reuse for industrial application has also been attempted.