Protein microbiology

Protein microbiology DEFAULT

Bacterial Protein Secretion


Protein export and secretion are fundamental processes for all forms of life. Secreted or membrane-associated proteins play a crucial role in many functions that are essential for viability and pathogenicity: cell structure maintenance, motility, cell attachment, metabolite transport, cell-cell interactions, and export of toxins. Bacteria transport these proteins out of their cytoplasm, where they are synthesized into their plasma membrane or across it, into the cell envelope or extracellular space. For all these protein export processes, bacteria have an array of various protein translocation pathways at their disposal, also depending on whether they are surrounded by a double or a single membrane bilayer.

The collection below highlights several of these transport systems, and our current understanding of their underlying molecular mechanisms. Deeper insight into these mechanisms provide a universal, fundamental understanding of protein trafficking mechanics, as many of the biochemical challenges that exported proteins encounter are shared across life. Such deeper understanding will enable the development of novel antibiotics and vaccines, the ability to combat pathogens through attenuation of virulence, and the improvement of strains and biomaterials for the production of biopharmaceuticals industrial enzymes, bioremediation, biofuel and chemicals production and various other industrial processes.

This FEMS Microbiology Letters Thematic Issue was Guest Edited by Jozef Anné, Lily Karamanou, and Tassos Economou to coincide with the FEMS sponsored meeting on Bacterial Protein Export 2018, Leuven, Belgium. This meeting will take place from 30 September – 3 October 2018.


Editorial: Thematic issue on bacterial protein export: from fundamentals to applications

Jozef Anné, Spyridoula Karamanou, Anastassios Economou

DOI: 10.1093/femsle/fny206


Bacterial machineries for the assembly of membrane-embedded β-barrel proteins

David Ranava, Anne Caumont-Sarcos, Cécile Albenne, Raffaele Ieva



Monitoring substrate enables real-time regulation of a protein localization pathway

Koreaki Ito, Hiroyuki Mori, Shinobu Chiba




YidC-mediated membrane insertion

Dorothee Kiefer, Andreas Kuhn





Structure-based working model of SecDF, a proton-driven bacterial protein translocation factor

Tomoya Tsukazaki




Hierarchical protein export mechanism of the bacterial flagellar type III protein export apparatus

Tohru Minamino




The Tat protein transport system: intriguing questions and conundrums

Shruthi Hamsanathan, Siegfried M Musser




Conformational plasticity of molecular chaperones involved in periplasmic and outer membrane protein folding

Guillaume Mas, Sebastian Hiller



Dynamic hydrogen-bond networks in bacterial protein secretion

Konstantina Karathanou, Ana-Nicoleta Bondar




SecA inhibitors as potential antimicrobial agents: differential actions on SecA-only and SecA-SecYEG protein-conducting channels

Jinshan Jin, Ying-Hsin Hsieh, Arpana S Chaudhary, Jianmei Cui, John E Houghton, Sen-fang Sui, Binghe Wang, Phang C Tai



Shaping Escherichia coli for recombinant membrane protein production

Alexandros Karyolaimos, Henry Ampah-Korsah, Zhe Zhang, Jan-Willem de Gier




Co-translational protein targeting in bacteria

Ruth Steinberg, Lara Knüpffer, Andrea Origi, Rossella Asti, Hans-Georg Koch




Type I secretion system—it takes three and a substrate

Kerstin Kanonenberg, Olivia Spitz, Isabelle N Erenburg, Tobias Beer, Lutz Schmitt



The way is the goal: how SecA transports proteins across the cytoplasmic membrane in bacteria

Tamar Cranford-Smith, Damon Huber




On display: autotransporter secretion and application

Peter van Ulsen, Katinka M Zinner, Wouter S P Jong, Joen Luirink




Bacterial secretion chaperones; the mycobacterial type VII case

Trang H Phan, Edith N G Houben




Bacterial type III secretion systems: A complex device for delivery of bacterial effector proteins into eukaryotic host cells

Samuel Wagner, Iwan Grin, Silke Malmsheimer, Nidhi Singh, Claudia E Torres-Vargas, Sibel Westerhausen



Streptomyces protein secretion and its application in biotechnology

Mohamed Belal Hamed, Jozef Anné, Spyridoula Karamanou, Anastassios Economou

DOI: 10.1093/femsle/fny250

Research Articles

Optimization of Type 3 protein secretion in enteropathogenic E.coli

Biao Yuan, Anastassios Economou, Spyridoula Karamanou



Recombinant protein expression in Escherichia coli: advances and challenges


There is no doubt that the production of recombinant proteins in microbial systems has revolutionized biochemistry. The days where kilograms of animal and plant tissues or large volumes of biological fluids were needed for the purification of small amounts of a given protein are almost gone. Every researcher that embarks on a new project that will need a purified protein immediately thinks of how to obtain it in a recombinant form. The ability to express and purify the desired recombinant protein in a large quantity allows for its biochemical characterization, its use in industrial processes and the development of commercial goods.

At the theoretical level, the steps needed for obtaining a recombinant protein are pretty straightforward. You take your gene of interest, clone it in whatever expression vector you have at your disposal, transform it into the host of choice, induce and then, the protein is ready for purification and characterization. In practice, however, dozens of things can go wrong. Poor growth of the host, inclusion body (IB) formation, protein inactivity, and even not obtaining any protein at all are some of the problems often found down the pipeline.

In the past, many reviews have covered this topic with great detail (Makrides, 1996; Baneyx, 1999; Stevens, 2000; Jana and Deb, 2005; Sorensen and Mortensen, 2005). Collectively, these papers gather more than 2000 citations. Yet, in the field of recombinant protein expression and purification, progress is continuously being made. For this reason, in this review, we comment on the most recent advances in the topic. But also, for those with modest experience in the production of heterologous proteins, we describe the many options and approaches that have been successful for expressing a great number of proteins over the last couple of decades, by answering the questions needed to be addressed at the beginning of the project. Finally, we provide a troubleshooting guide that will come in handy when dealing with difficult-to-express proteins.

First Question: Which Organism to Use?

The choice of the host cell whose protein synthesis machinery will produce the precious protein will initiate the outline of the whole process. It defines the technology needed for the project, be it a variety of molecular tools, equipment, or reagents. Among microorganisms, host systems that are available include bacteria, yeast, filamentous fungi, and unicellular algae. All have strengths and weaknesses and their choice may be subject to the protein of interest (Demain and Vaishnav, 2009; Adrio and Demain, 2010). For example, if eukaryotic post-translational modifications (like protein glycosylation) are needed, a prokaryotic expression system may not be suitable (Sahdev et al., 2008). In this review, we will focus specifically on Escherichia coli. Other systems are described in excellent detail in accompanying articles of this series.

The advantages of using E. coli as the host organism are well known. (i) It has unparalleled fast growth kinetics. In glucose-salts media and given the optimal environmental conditions, its doubling time is about 20 min (Sezonov et al., 2007). This means that a culture inoculated with a 1/100 dilution of a saturated starter culture may reach stationary phase in a few hours. However, it should be noted that the expression of a recombinant protein may impart a metabolic burden on the microorganism, causing a considerable decrease in generation time (Bentley et al., 1990). (ii) High cell density cultures are easily achieved. The theoretical density limit of an E. coli liquid culture is estimated to be about 200 g dry cell weight/l or roughly 1 × 1013 viable bacteria/ml (Lee, 1996; Shiloach and Fass, 2005). However, exponential growth in complex media leads to densities nowhere near that number. In the simplest laboratory setup (i.e., batch cultivation of E. coli at 37°C, using LB media), <1 × 1010 cells/ml may be the upper limit (Sezonov et al., 2007), which is less than 0.1% of the theoretical limit. For this reason, high cell-density culture methods were designed to boost E. coli growth, even when producing a recombinant protein (Choi et al., 2006). Being a workhorse organism, these strategies arose thanks to the wealth of knowledge about its physiology. (iii) Rich complex media can be made from readily available and inexpensive components. (iv) Transformation with exogenous DNA is fast and easy. Plasmid transformation of E. coli can be performed in as little as 5 min (Pope and Kent, 1996).

Second Question: Which Plasmid Should Be Chosen?

The most common expression plasmids in use today are the result of multiple combinations of replicons, promoters, selection markers, multiple cloning sites, and fusion protein/fusion protein removal strategies (Figure 1). For this reason, the catalog of available expression vectors is huge and it is easy to get lost when choosing a suitable one. To make an informed decision, these features have to be carefully evaluated according to the individual needs.

FIGURE 1. Anatomy of an expression vector. The figure depicts the major features present in common expression vectors. All of them are described in the text. The affinity tags and coding sequences for their removal were positioned arbitrarily at the N-terminus for simplicity. MCS, multiple cloning site. Striped patterned box: coding sequence for the desired protein.


Genetic elements that undergo replication as autonomous units, such as plasmids, contain a replicon. It consists of one origin of replication together with its associated cis-acting control elements. An important parameter to have in mind when choosing a suitable vector is copy number. The control of copy number resides in the replicon (del Solar and Espinosa, 2000). It is logical to think that high plasmid dosage equals more recombinant protein yield as many expression units reside in the cell. However, a high plasmid number may impose a metabolic burden that decreases the bacterial growth rate and may produce plasmid instability, and so the number of healthy organisms for protein synthesis falls (Bentley et al., 1990; Birnbaum and Bailey, 1991). For this reason, the use of high copy number plasmids for protein expression by no means implies an increase in production yields.

Commonly used vectors, such as the pET series, possess the pMB1 origin (ColE1-derivative, 15–60 copies per cell; Bolivar et al., 1977) while a mutated version of the pMB1 origin is present in the pUC series (500–700 copies per cell; Minton, 1984). The wild-type ColE1 origin (15–20 copies per cell; Lin-Chao and Bremer, 1986; Lee et al., 2006) can be found in the pQE vectors (Qiagen). They all belong to the same incompatibility group meaning that they cannot be propagated together in the same cell as they compete with each other for the replication machinery (del Solar et al., 1998; Camps, 2010). For the dual expression of recombinant proteins using two plasmids, systems with the p15A ori are available (pACYC and pBAD series of plasmids, 10–12 copies per cell; Chang and Cohen, 1978; Guzman et al., 1995). Though rare, triple expression can be achieved by the use of the pSC101 plasmid. This plasmid is under a stringent control of replication, thus it is present in a low copy number (<5 copies per cell; Nordstrom, 2006). The use of plasmids bearing this replicon can be an advantage in cases where the presence of a high dose of a cloned gene or its product produces a deleterious effect to the cell (Stoker et al., 1982; Wang and Kushner, 1991). Alternatively, the use of the Duet vectors (Novagen) simplifies dual expression by allowing cloning of two genes in the same plasmid. The Duet plasmids possess two multiple cloning sites, each preceded by a T7 promoter, a lac operator and a ribosome binding site. By combining different compatible Duet vectors, up to eight recombinant proteins can be produced from four expression plasmids.


The staple in prokaryotic promoter research is undoubtedly the lac promoter, key component of the lac operon (Müller-Hill, 1996). The accumulated knowledge in the functioning of the system allowed for its extended use in expression vectors. Lactose causes induction of the system and this sugar can be used for protein production. However, induction is difficult in the presence of readily metabolizable carbon sources (such as glucose present in rich media). If lactose and glucose are present, expression from the lac promoter is not fully induced until all the glucose has been utilized. At this point (low glucose), cyclic adenosine monophosphate (cAMP) is produced, which is necessary for complete activation of the lac operon (Wanner et al., 1978; Postma and Lengeler, 1985). This positive control of expression is known as catabolite repression. In accordance, cAMP levels are low in cells growing in lac operon-repressing sugars, and this correlates with lower rates of expression of the lac operon (Epstein et al., 1975). Also, glucose abolishes lactose uptake because lactose permease is inactive in the presence of glucose (Winkler and Wilson, 1967). To achieve expression in the presence of glucose, a mutant that reduces (but does not eliminate) sensitivity to catabolite regulation was introduced, the lacUV5 promoter (Silverstone et al., 1970; Lanzer and Bujard, 1988). However, when present in multicopy plasmids, both promoters suffer from the disadvantage of sometimes having unacceptably high levels of expression in the absence of inducer (a.k.a. “leakiness”) due to titration of the low levels of the lac promoter repressor protein LacI from the single chromosomal copy of its gene (about 10 molecules per cell; Müller-Hill et al., 1968). Basal expression control can be achieved by the introduction of a mutated promoter of the lacI gene, called lacIQ, that leads to higher levels of expression (almost 10-fold) of LacI (Calos, 1978). The lac promoter and its derivative lacUV5 are rather weak and thus not very useful for recombinant protein production (Deuschle et al., 1986; Makoff and Oxer, 1991). Synthetic hybrids that combine the strength of other promoters and the advantages of the lac promoter are available. For example, the tac promoter consists of the -35 region of the trp (tryptophan) promoter and the -10 region of the lac promoter. This promoter is approximately 10 times stronger than lacUV5 (de Boer et al., 1983). Notable examples of commercial plasmids that use the lac or tac promoters to drive protein expression are the pUC series (lacUV5 promoter, Thermo Scientific) and the pMAL series of vectors (tac promoter, NEB).

The T7 promoter system present in the pET vectors (pMB1 ori, medium copy number, Novagen) is extremely popular for recombinant protein expression. This is not surprising as the target protein can represent 50% of the total cell protein in successful cases (Baneyx, 1999; Graumann and Premstaller, 2006). In this system, the gene of interest is cloned behind a promoter recognized by the phage T7 RNA polymerase (T7 RNAP). This highly active polymerase should be provided in another plasmid or, most commonly, it is placed in the bacterial genome in a prophage (λDE3) encoding for the T7 RNAP under the transcriptional control of a lacUV5 promoter (Studier and Moffatt, 1986). Thus, the system can be induced by lactose or its non-hydrolyzable analog isopropyl β-D-1-thiogalactopyranoside (IPTG). Basal expression can be controlled by lacIQ but also by T7 lysozyme co-expression (Moffatt and Studier, 1987). T7 lysozyme binds to T7 RNAP and inhibits transcription initiation from the T7 promoter (Stano and Patel, 2004). In this way, if small amounts of T7 RNAP are produced because of leaky expression of its gene, T7 lysozyme will effectively control unintended expression of heterologous genes placed under the T7 promoter. T7 lysozyme is provided by a compatible plasmid (pLysS or pLysE). After induction, the amount of T7 RNAP produced surpasses the level of polymerase that T7 lysozyme can inhibit. The “free” T7 RNAP can thus engage in transcription of the recombinant gene. Yet another level of control lies in the insertion of a lacO operator downstream of the T7 promoter, making a hybrid T7/lac promoter (Dubendorff and Studier, 1991). All three mechanisms (tight repression of the lac-inducible T7 RNAP gene by lacIQ, T7 RNAP inhibition by T7 lysozyme and presence of a lacO operator after the T7 promoter) make the system ideal for avoiding basal expression.

The problem of leaky expression is a reflection of the negative control of the lac promoter. Promoters that rely on positive control should have lower background expression levels (Siegele and Hu, 1997). This is the case of the araPBAD promoter present in the pBAD vectors (Guzman et al., 1995). The AraC protein has the dual role of repressor/activator. In the absence of arabinose inducer, AraC represses translation by binding to two sites in the bacterial DNA. The protein–DNA complex forms a loop, effectively preventing RNA polymerase from binding to the promoter. Upon addition of the inducer, AraC switches into “activation mode” and promotes transcription from the ara promoter (Schleif, 2000, 2010). In this way, arabinose is absolutely needed for induction.

Another widely used approach is to place a gene under the control of a regulated phage promoter. The strong leftward promoter (pL) of phage lambda directs expression of early lytic genes (Dodd et al., 2005). The promoter is tightly repressed by the λcI repressor protein, which sits on the operator sequences during lysogenic growth. When the host SOS response is triggered by DNA damage, the expression of the protein RecA is stimulated, which in turn catalyzes the self-cleavage of λcI, allowing transcription of pL-controlled genes (Johnson et al., 1981; Galkin et al., 2009). This mechanism is used in expression vectors containing the pL promoter. The SOS response (and recombinant protein expression) can be elicited by adding nalidixic acid, a DNA gyrase inhibitor (Lewin et al., 1989; Shatzman et al., 2001). Another way of activating the promoter is to control λcI production by placing its gene under the influence of another promoter. This two-stage control system has already been described for T7 promoter/T7 RNAP-based vectors. In the pLEX series of vectors (Life Technologies), the λcI repressor gene was integrated into the bacterial chromosome under the control of the trp promoter. In the absence of tryptophan, this promoter is always “on” and λcI is continuously produced. Upon addition of tryptophan, a tryptophan-TrpR repressor complex is formed that tightly binds to the trp operator, thereby blocking λcI repressor synthesis. Subsequently, the expression of the desired gene under the pL promoter ensues (Mieschendahl et al., 1986).

Transcription from all promoters discussed so far is initiated by chemical cues. Systems that respond to physical signals (e.g., temperature or pH) are also available (Goldstein and Doi, 1995). The pL promoter is one example. A mutant λcI repressor protein ( λcI857) is temperature-sensitive and is unstable at temperatures higher than 37°C. E. coli host strains containing the λcI857 protein (either integrated in the chromosome or into a vector) are first grown at 28–30°C to the desired density, and then protein expression is induced by a temperature shift to 40–42°C (Menart et al., 2003; Valdez-Cruz et al., 2010). The industrial advantage of this system lies in part in the fact that during fermentation, heat is usually produced and increasing the temperature in high density cultures is easy. On the other hand, genes under the control of the cold-inducible promoter cspA are induced by a downshift in temperature to 15°C (Vasina et al., 1998). This temperature is ideal for expressing difficult proteins as will be explained in another section. The pCold series of plasmids have a pUC118 backbone (a pUC18 derivative; Vieira and Messing, 1987) with the cspA promoter (Qing et al., 2004; Hayashi and Kojima, 2008). In the original paper, successful expression was achieved for more than 30 recombinant proteins from different sources, reaching levels as high as 20–40% of the total expressed proteins (Qing et al., 2004). However, it should be noted that in various cases the target proteins were obtained in an insoluble form.

Selection Marker

To deter the growth of plasmid-free cells, a resistance marker is added to the plasmid backbone. In the E. coli system, antibiotic resistance genes are habitually used for this purpose. Resistance to ampicillin is conferred by the bla gene whose product is a periplasmic enzyme that inactivates the β-lactam ring of β-lactam antibiotics. However, as the β-lactamase is continuously secreted, degradation of the antibiotic ensues and in a couple of hours, ampicillin is almost depleted (Korpimaki et al., 2003). Under this situation, cells not carrying the plasmid are allowed to increase in number during cultivation. Although not experimentally verified, selective agents in which resistance is based on degradation, like chloramphenicol (Shaw, 1983) and kanamycin (Umezawa, 1979), could also have this problem. For this reason, tetracycline has been shown to be highly stable during cultivation (Korpimaki et al., 2003), because resistance is based on active efflux of the antibiotic from resistant cells (Roberts, 1996).

The cost of antibiotics and the dissemination of antibiotic resistance are major concerns in projects dealing with large-scale cultures. Much effort has been put in the development of antibiotics-free plasmid systems. These systems are based on the concept of plasmid addiction, a phenomenon that occurs when plasmid-free cells are not able to grow or live (Zielenkiewicz and Ceglowski, 2001; Peubez et al., 2010). For example, an essential gene can be deleted from the bacterial genome and then placed on a plasmid. Thus, after cell division, plasmid-free bacteria die. Different subtypes of plasmid-addiction systems exist according to their principle of function: (i) toxin/antitoxin-based systems, (ii) metabolism-based systems, and (iii) operator repressor titration systems (Kroll et al., 2010). While this promising technology has been proved successful in large-scale fermentors (Voss and Steinbuchel, 2006; Peubez et al., 2010), expression systems based on plasmid addiction are still not widely distributed.

Affinity Tags

When devising a project where a purified soluble active recombinant protein is needed (as is often the case), it is invaluable to have means to (i) detect it along the expression and purification scheme, (ii) attain maximal solubility, and (iii) easily purify it from the E. coli cellular milieu. The expression of a stretch of amino acids (peptide tag) or a large polypeptide (fusion partner) in tandem with the desired protein to form a chimeric protein may allow these three goals to be straightforwardly reached (Nilsson et al., 1997).

Being small, peptide tags are less likely to interfere when fused to the protein. However, in some cases they may provoke negative effects on the tertiary structure or biological activity of the fused chimeric protein (Bucher et al., 2002; Klose et al., 2004; Chant et al., 2005; Khan et al., 2012). Vectors are available that allow positioning of the tag on either the N-terminal or the C-terminal end (the latter option being advantageous when a signal peptide is positioned at the N-terminal end for secretion of the recombinant protein, see below). If the three-dimensional structure of the desired protein is available, it is wise to check which end is buried inside the fold and place the tag in the solvent-accessible end. Common examples of small peptide tags are the poly-Arg-, FLAG-, poly-His-, c-Myc-, S-, and Strep II-tags (Terpe, 2003). Since commercial antibodies are available for all of them, the tagged recombinant protein can be detected by Western blot along expression trials, which is extremely helpful when the levels of the desired proteins are not high enough to be detected by SDS-PAGE. Also, tags allow for one-step affinity purification, as resins that tightly and specifically bind the tags are available. For example, His-tagged proteins can be recovered by immobilized metal ion affinity chromatography using Ni2+ or Co2+-loaded nitrilotriacetic acid-agarose resins (Porath and Olin, 1983; Bornhorst and Falke, 2000), while anti-FLAG affinity gels (Sigma-Aldrich) are used for capturing FLAG fusion proteins (Hopp et al., 1988).

On the other hand, adding a non-peptide fusion partner has the extra advantage of working as solubility enhancers (Hammarstrom et al., 2002). The most popular fusion tags are the maltose-binding protein (MBP; Kapust and Waugh, 1999), N-utilization substance protein A (NusA; Davis et al., 1999), thioredoxin (Trx; LaVallie et al., 1993), glutathione S-transferase (GST; Smith and Johnson, 1988), ubiquitin (Baker, 1996) and SUMO (Butt et al., 2005). The reasons why these fusion partners act as solubility enhancers remain unclear and several hypothesis have been proposed (reviewed in Raran-Kurussi and Waugh, 2012). In the case of MBP, it was shown that it possesses an intrinsic chaperone activity (Kapust and Waugh, 1999; Raran-Kurussi and Waugh, 2012). In comparison studies, GST showed the poorest solubility enhancement capabilities (Hammarstrom et al., 2006; Bird, 2011). NusA, MBP, and Trx display the best solubility enhancing properties but their large size may lead to the erroneous assessment of protein solubility (Costa et al., 2013). Indeed, when these tags are removed, the final solubility of the desired product is unpredictable (Esposito and Chatterjee, 2006). For these reasons, smaller tags with strong solubility enhancing effects are desirable. Recently, the 8-kDa calcium binding protein Fh8 from the parasite Fasciola hepatica was shown to be as good as or better than the large tags in terms of solubility enhancement. Moreover, the recombinant proteins maintained their solubility after tag removal (Costa et al., 2013). MBP and GST can be used to purify the fused protein by affinity chromatography, as MBP binds to amylose–agarose and GST to glutathione–agarose. MBP is present in the pMAL series of vectors from NEB and GST in the pGEX series (GE). A peptide tag must be added to the fusion partner-containing protein if an affinity chromatography step is needed in the purification scheme. MBP and GST bind to their substrates non-covalently. On the contrary, the HaloTag7 (Promega) is based on the covalent capture of the tag to the resin, making the system fast and highly specific (Ohana et al., 2009).

A different group of fusion tags are stimulus-responsive tags, which reversibly precipitate out of solution when subjected to the proper stimulus. The addition of β roll tags to a recombinant protein allows for its selective precipitation in the presence of calcium. The final products presented a high purity and the precipitation protocol only takes a couple of minutes (Shur et al., 2013). Another protein-based stimulus-responsive purification tags are elastin-like polypeptides (ELPs), which consist of tandem repeats of the sequence VPGXG, where X is Val, Ala, or Gly in a 5:2:3 ratio (Meyer and Chilkoti, 1999). These tags undergo an inverse phase transition at a given temperature of transition (Tt). When the Tt is reached, the ELP–protein fusion selectively and reversibly precipitates, allowing for quick enrichment of the recombinant protein by centrifugation (Banki et al., 2005). Precipitation can also be triggered by adjusting the ionic strength of the solution (Ge et al., 2005). These techniques represent an alternative to conventional chromatography-based purification methods and can save production costs, especially in large-scale settings (Fong and Wood, 2010). The main characteristics of the tags mentioned in this section are outlined on Table 1.

TABLE 1. Main characteristics of protein fusion tags.

Tag Removal

If structural or biochemical studies on the recombinant protein are needed, then the fusion partner must be eliminated from the recombinant protein. Peptide tags should be removed too because they can interfere with protein activity and structure (Wu and Filutowicz, 1999; Perron-Savard et al., 2005), but they can be left in place even for crystallographic studies (Bucher et al., 2002; Carson et al., 2007). Tags can be eliminated by either enzymatic cleavage or chemical cleavage.

In the case of tag removal by enzyme digestion, expression vectors possess sequences that encode for protease cleavage sites downstream of the gene coding for the tag. Enterokinase, thrombin, factor Xa and the tobacco etch virus (TEV) protease have all been successfully used for the removal of peptide tags and fusion partners (Jenny et al., 2003; Blommel and Fox, 2007). Choosing among the different proteases is based on specificity, cost, number of amino acids left in the protein after cleavage and ease of removal after digestion (Waugh, 2011). Enterokinase and thrombin were popular in the past but the use of His-tagged TEV has become an everyday choice due to its high specificity (Parks et al., 1994), it is easy to produce in large quantities (Tropea et al., 2009) and leaves only a serine or glycine residue (or even the natural N-terminus) after digestion (Kapust et al., 2002).

As the name implies, in chemical cleavage the tag is removed by treatment of the fusion protein with a chemical reagent. The advantages of using chemicals for this purpose are that they are easy to eliminate from the reaction mixture and are cheap in comparison with proteolytic enzymes, which makes them an attractive choice in the large-scale production of recombinant proteins (Rais-Beghdadi et al., 1998). However, the reaction conditions are harsh, so their use is largely restricted to purified recombinant proteins obtained from IBs. They also often cause unwanted protein modifications (Hwang et al., 2014). The most common chemical cleavage reagent is cyanogen bromide (CNBr). CNBr cleaves the peptide bond C-terminal to methionine residues, so this amino acid should be present between the tag and the protein of interest (Rais-Beghdadi et al., 1998). Also, the target protein should not contain internal methionines. CNBr cleavage can be performed in common denaturing conditions (6 M guanidinium chloride) or 70% formic acid or trifluoroacetic acid (Andreev et al., 2010). Other chemical methods for protein cleavage can be found in Hwang et al. (2014).

Third Question: Which is the Appropriate Host?

A quick search in the literature for a suitable E. coli strain to use as a host will yield dozens of possible candidates. All of them have advantages and disadvantages. However, something to keep in mind is that many are specialty strains that are used in specific situations. For a first expression screen, only a couple of E. coli strains are necessary: BL21(DE3) and some derivatives of the K-12 lineage.

The history of the BL21 and BL21(DE3) strains was beautifully documented in Daegelen et al. (2009) and we recommend this article to the curious. BL21 was described by Studier in 1986 after various modifications of the B line (Studier and Moffatt, 1986), which in turn Daegelen et al. (2009) traced back to d’Herelle. A couple of genetic characteristics of BL21 are worthy of mention. Like other parental B strains, BL21 cells are deficient in the Lon protease, which degrades many foreign proteins (Gottesman, 1996). Another gene missing from the genome of the ancestors of BL21 is the one coding for the outer membrane protease OmpT, whose function is to degrade extracellular proteins. The liberated amino acids are then taken up by the cell. This is problematic in the expression of a recombinant protein as, after cell lysis, OmpT may digest it (Grodberg and Dunn, 1988). In addition, plasmid loss is prevented thanks to the hsdSB mutation already present in the parental strain (B834) that gave rise to BL21. As a result, DNA methylation and degradation is disrupted. When the gene of interest is placed under a T7 promoter, then T7 RNAP should be provided. In the popular BL21(DE3) strain, the λDE3 prophage was inserted in the chromosome of BL21 and contains the T7 RNAP gene under the lacUV5 promoter, as was explained earlier.

The BL21(DE3) and its derivatives are by far the most used strains for protein expression. Still, there are reports where the K-12 lineage is used for this purpose. The AD494 and OrigamiTM (Novagen) strains are trxB (thioredoxin reductase) mutants, so disulfide bond formation in the cytoplasm is enhanced (the Origami strain also lacks the glutathione reductase gene; Derman et al., 1993). Another widely used strain from the K-12 repertoire is HMS174, a recA mutant (Campbell et al., 1978). This mutation has a positive effect on plasmid stability (Marisch et al., 2013). Plasmid multimer formation, an important cause of instability, relies on the recombination system of E. coli (Summers et al., 1993). All three strains have their λDE3-containing derivative (available at Novagen) so the T7 RNAP system can be used.

Fourth Question: Which is the Combination for Success?

At this point, it should be pretty clear that the number of options when designing an expression system is considerably high. Choosing the perfect combination is not possible a priori, so multiple conditions should be tested to obtain the desired protein. If the project demands expressing two protein constructs, cloned in six different expression vectors, each transformed in three different expression strains, then you are in for 36 expression trials. This number may be even higher when other variables are taken into account. This trial-and-error and time consuming pilot study can be made faster if micro-expression trials are performed before scale-up. Small-scale screens can be performed in 2-ml tubes or 96-well plates (Shih et al., 2002). High throughput protocols adapting automatic liquid handling robots have been described, making it possible for a single person to test more than 1000 culture conditions within a week.

Troubleshooting Recombinant Protein Production

This section of the review covers different strategies for optimizing recombinant protein production in E. coli. Even after careful selection of plasmid and host, it cannot be predicted if the protein will be obtained in high amounts and in a soluble active form. Various situations that impede reaching that goal can be encountered, which unfortunately happen very often. Many things to try in each case are discussed in the following paragraphs and, for convenience of the readers; a summary is included in Table 2.

TABLE 2. Strategies for overcoming common problems during recombinant protein expression in E. coli.

No or Low Production

This situation may be regarded as the worst case scenario. When the protein of interest cannot be detected through a sensitive technique (e.g., Western blot) or it is detected but at very low levels (less than micrograms per liter of culture), the problem often lies in a harmful effect that the heterologous protein exerts on the cell (Miroux and Walker, 1996; Dumon-Seignovert et al., 2004).

Protein toxicity

The problem of protein toxicity may arise when the recombinant protein performs an unnecessary and detrimental function in the host cell. This function interferes with the normal proliferation and homeostasis of the microorganism and the visible result is slower growth rate, low final cell density, and death (Doherty et al., 1993; Dong et al., 1995).

As a first measure, cell growth should be monitored before induction. If the growth rate of the recombinant strain is slower compared to an empty-vector bearing strain then two causes may explain the phenotype: gene toxicity and basal expression of the toxic mRNA/protein. Gene toxicity will not be discussed here and the review of Saida et al. (2006) is recommended.

The control of basal synthesis was covered in some detail in Section “Promoter.” As stated, the expression of LacI from lacI or lacIQ represses transcription of lac-based promoters. For high copy number plasmids (>100 copies per cell), lacIQ should be cloned in the expression vector. The pQE vectors from Qiagen utilize two lac operator sequences to increase control of the T5 promoter, which is recognized by the E. coli RNA polymerase (see The QIAexpressionistTM manual from Qiagen). A tighter control can be achieved by the addition of 0.2–1% w/v glucose in the medium as rich media prepared with tryptone or peptone may contain the inducer lactose (Studier, 2005). Another option could be to prepare defined media using glucose as a source of carbon. In T7-based promoters, leaky expression is avoided by co-expression of T7 lysozyme from the pLysS or pLysE plasmids (see above). Use of lower copy number plasmids containing tightly regulated promoters (like the araPBAD promoter) is suggested. An interesting case of copy number control is the one employed in pETcoco vectors (Novagen). These plasmids possess two origins of replication. The oriS origin and its control elements maintain pETcoco at one copy per cell (Wild et al., 2002). However, the TrfA replicator activates the medium-copy origin of replication (oriV) and amplification of copy number is achieved (up to 40 copies per cell). The trfA gene is on the same vector and is under control of the araPBAD promoter, so copy number can be controlled by arabinose (Wild et al., 2002).

After control of basal expression, the culture should grow well until the proper time of induction. At this moment, if the protein is toxic, cell growth will be arrested. In many cases, the level of toxicity of a protein becomes apparent when a certain threshold of host tolerance is reached and exceeded. In such situations, the level of expression should be manipulated at will. Tunable expression can be achieved using the Lemo21(DE3) strain. This strain is similar to the BL21(DE3)pLysS strain, however, T7 lysozyme production from the lysY gene is under the tunable promoter rhaPBAD (Wagner et al., 2008). At higher concentrations of the sugar L-rhamnose, more T7 lysozyme is produced, less active T7 RNAP is present in the cell and less recombinant protein is expressed. Trials using L-rhamnose concentrations from 0 to 2,000 μM should be undertaken to find the best conditions for expression. By contrast, dose-dependent expression when using IPTG as inducer is not possible since IPTG can enter the cell by active transport through the Lac permease or by permease-independent pathways (Fernandez-Castane et al., 2012). Since expression of Lac permease is heterogeneous and the number of active permeases in each cell is highly variable, protein expression does not respond predictably to IPTG concentration. The TunerTM (DE3) strain (Novagen) is a BL21 derivative that possesses a lac permease (lacY) mutation that allows uniform entry of IPTG into all LacY- cells in the population, which produces a concentration-dependent, homogeneous level of induction (Khlebnikov and Keasling, 2002). In the same line of thought, an E. coli strain was constructed by exchanging the wild-type operator by the derivative lacOc, thus converting the lac operon into a constitutive one. This modification avoids the transient non-genetic LacY- phenotype of a fraction of the cells, allowing uniform entry of the inducer lactose. A second modification (gal+) permits the full utilization of lactose as an energy source (Menzella et al., 2003).

A word of caution needs to be said in regard to “tunable promoters” that are inducible by sugars (lactose, arabinose, rhamnose). In the case of the araPBAD promoter, the yields of the target protein can be reproducibly increased over a greater than 100-fold range by supplementing the culture with different sub-maximal concentrations of arabinose (Guzman et al., 1995). This led to the erroneous belief that within each cell, the level of recombinant protein synthesis can be manipulated at will. However, it was shown that the range in protein expression arises from the heterogeneity in the amount of active sugar permeases in each cell, as was also explained for LacY (Siegele and Hu, 1997). So, even though the final protein yield can be controlled, the amount of protein per cell is widely variable, with cells producing massive amounts of protein and others not producing any protein at all. This can be a nuance, since in the case of toxic products; the subpopulation of cells with high-level synthesis may perish (Doherty et al., 1993; Dong et al., 1995).

Some E. coli mutants were specifically selected to withstand the expression of toxic proteins. The strains C41(DE3) and C43(DE3) were found by Miroux and Walker (1996) in a screen designed to isolate derivatives of BL21(DE3) with improved membrane protein overproduction characteristics. It was recently discovered that the previously uncharacterized mutations which prevent cell death during the expression of recombinant proteins in these strains lie on the lacUV5 promoter. In BL21(DE3) cells, the lacUV5 promoter drives the expression of the T7 RNAP, but in the Walker strains two mutations in the -10 region revert the lacUV5 promoter back into the weaker wild-type counterpart. This leads to a lesser (and perhaps more tolerable for the cell) level of synthesis (Wagner et al., 2008).

Another solution could be to remove the protein from the cell. Secretion to the periplasm or to the medium is sometimes the only way to produce a recombinant protein (Mergulhao et al., 2005; de Marco, 2009). The first option for expression in the periplasm is the post-translational Sec-dependent pathway (Georgiou and Segatori, 2005). Routing to the extracytoplasmatic space is achieved by fusing the recombinant protein to a proper leader peptide. The signal peptides of the following proteins are widely used for secretion: Lpp, LamB, LTB, MalE, OmpA, OmpC, OmpF, OmpT, PelB, PhoA, PhoE, or SpA (Choi and Lee, 2004). The co-translational translocation machinery based on the SRP (signal recognition particle) pathway can also be used. SRP recognizes its substrates by the presence of a hydrophobic signal sequence located in the N-terminal end. Following interaction with the membrane receptor FtsY, the complex of nascent chain and ribosome is transferred to the SecYEG translocase (Valent et al., 1998). The signal sequence of disulfide isomerase I (DsbA) has been used to target recombinant proteins to the periplasm via the SRP pathway. Notable examples of recombinant proteins secreted though this system include thioredoxin (Schierle et al., 2003) and the human growth hormone (Soares et al., 2003).

Codon bias

Codon bias arises when the frequency of occurrence of synonymous codons in the foreign coding DNA is significantly different from that of the host. At the moment of full synthesis of the recombinant protein, depletion of low-abundance tRNAs occurs. This deficiency may lead to amino acid misincorporation and/or truncation of the polypeptide, thus affecting the heterologous protein expression levels (which will be low at best) and/or its activity (Gustafsson et al., 2004). To check if codon bias could be an issue when expressing a recombinant protein, a large number of free online apps detect the presence of rare codons in a given gene when E. coli is used as a host (,,, just to name a few). Rare codons were defined as codons used by E. coli at a frequency <1% (Kane, 1995). For example, the AGG codon (Arg) is used in E. coli at a frequency of <0.2%, but it is not rare in plant mRNAs where it can reach frequencies >1.5%.

Two strategies for solving codon usage bias have been used: codon optimization of the foreign coding sequence or increasing the availability of underrepresented tRNAs by host modification (Sorensen and Mortensen, 2005). The rationale behind codon usage optimization is to modify the rare codons in the target gene to mirror the codon usage of the host (Burgess-Brown et al., 2008; Welch et al., 2009; Menzella, 2011). The amino acid sequence of the encoded protein must not be altered in the process. This can be done by site-directed silent mutagenesis or resynthesis of the whole gene or parts of it. Codon optimization by silent mutagenesis is a cumbersome and expensive process, so is not very useful when many recombinant proteins are needed. On the other hand, gene synthesis by design is not a trivial issue since it requires choosing the best sequence from a vast number of possible combinations (Gustafsson et al., 2004). The simplest approach is to replace all instances of a given amino acid in the target gene by the most abundant codon of the host, a strategy called “one amino acid-one codon.” More advanced algorithms, which employ several other optimization parameters such as codon context and codon harmonization, have been described (Gao et al., 2004; Supek and Vlahovicek, 2004; Jayaraj et al., 2005; Angov et al., 2011). Some are freely available as web servers or standalone software. For a comprehensive list, please refer to Puigbo et al. (2007).

Correcting codon usage is a tricky situation. The “one amino acid-one codon” strategy disregards factors other than codon rarity that influence protein expression levels. For example, in bacterial genes enriched in rare codons at the N-terminus, protein expression is actually improved. The cause lies not in codon rarity per se but in the reduction of RNA secondary structure (Goodman et al., 2013). In addition, a recent report has shown that high levels of protein production are mainly (but not only) determined by the decoding speed of the open reading frame (i.e., the time it takes for a ribosome to translate an mRNA), especially if “fast” codons are located at the 5′-end of the mRNA (Chu et al., 2014). This causes a fast ribosome clearance at the initiation site, so that new recruited ribosomes encounter a free start codon and can engage in translation. Finally, some codon combinations can create Shine–Dalgarno-like structures that cause translational pausing by hybridization between the target mRNA and the 16S rRNA of the translating ribosome (Li et al., 2012). Translational pausing along the mRNA has a beneficial effect in protein folding, as it allows for the newly synthesized chain to adopt a well-folded intermediate conformation (Thanaraj and Argos, 1996; Oresic and Shalloway, 1998; Tsai et al., 2008; Yona et al., 2013). All of this new evidence in translational control mechanisms poses a challenge in the rational design of synthetic genes. Newer algorithms should account for 5′ RNA structure, presence of strategically located Shine–Dalgarno-like motifs, ribosome clearance rates at the initiation site and presence of slowly translated regions that are beneficial in co-translational folding.

On the other hand, when the cell is producing massive amounts of proteins (as in the case of recombinant expression of heterologous genes), charged tRNA availability for rare codons does become the major determinant of the levels of produced protein (Pedersen, 1984; Li et al., 2012). Low-abundance tRNA depletion causes ribosome stalling and its subsequent detachment from the RNA strand and thus, failure to generate a full-length product (Buchan and Stansfield, 2007). Several strains carrying plasmids containing extra copies of problematic tRNAs genes can be used to circumvent this issue. The BL21(DE3)CodonPlus strain (Stratagene) contains the pRIL plasmid (p15A replicon, which is compatible with the ColE1 and ColE1-like origins contained in most commonly used expression vectors), which provides extra genes for the tRNAs for AGG/AGA (Arg), AUA (Ile), and CUA (Leu). BL21(DE3)CodonPlus-RP (Stratagene) corrects for the use of AGG/AGA (Arg) and CCC (Pro). The Rosetta(DE3) strains (Novagen) are TunerTM derivatives containing the pRARE plasmid (p15A replicon), supplying tRNAs for all the above-mentioned codons plus GGA (Gly). It should be noted that the use of these strains often improves the levels of protein production but sometimes can cause a decrease in protein solubility. We have found that proteins with higher than 5% content of RIL codons (AGG/AGA, AUA, and CUA) are less soluble when expressed in the CodonPlus strain. In this host, the translational pauses introduced by the RIL codons are probably overridden, increasing translation speed and consequently, protein aggregation (Rosano and Ceccarelli, 2009).

Limiting factors in batch cultivation

When the expression of the recombinant protein is low and cannot be increased by the proposed mechanisms, then the volumetric yield of desired protein can be augmented by growing the culture to higher densities. This can be achieved by changing a few parameters, like medium composition and providing better aeration by vigorous shaking (McDaniel and Bailey, 1969; Cui et al., 2006; Blommel et al., 2007).

LB is the most commonly used medium for culturing E. coli. It is easy to make, it has rich nutrient contents and its osmolarity is optimal for growth at early log phase. All these features make it adequate for protein production and compensate for the fact that it is not the best option for achieving high cell density cultures. Despite being a rich broth, cell growth stops at a relatively low density. This happens because LB contains scarce amounts of carbohydrates (and other utilizable carbon sources) and divalent cations (Sezonov et al., 2007). Not surprisingly, increasing the amount of peptone or yeast extract leads to higher cell densities (Studier, 2005). Also, divalent cation supplementation (MgSO4 in the millimolar range) results in higher cell growth. Adding glucose is of limited help in this regard because acid generation by glucose metabolism overwhelms the limited buffer capacity of LB, at least in shake flasks where pH control can be laborious (Weuster-Botz et al., 2001; Scheidle et al., 2011). If culture acidification poses a problem, the media can be buffered with phosphate salts at 50 mM. 2xYT, TB (Terrific Broth) and SB (Super Broth) media recipes are available elsewhere and have been shown to be superior to LB for reaching higher cell densities (Madurawe et al., 2000; Atlas, 2004; Studier, 2005).

A major breakthrough in media composition came in 2005 by the extensive work of Studier. In that report, the concept of autoinduction was developed (Studier, 2005). In autoinduction media, a mixture of glucose, lactose, and glycerol is used in an optimized blend. Glucose is the preferred carbon source and is metabolized preferentially during growth, which prevents uptake of lactose until glucose is depleted, usually in mid to late log phase. Consumption of glycerol and lactose follows, the latter being also the inducer of lac-controlled protein expression. In this way, biomass monitoring for timely inducer addition is avoided, as well as culture manipulation (Studier, 2014).

As the number of cells per liter increases, oxygen availability becomes an important factor with profound influence on growth (O’Beirne and Hamer, 2000; Losen et al., 2004).Oxygen limitation triggers the expression of more than 200 genes in an attempt to adjust the metabolic capacities of the cell to the availability of oxygen, all of which hinder optimal growth over long culture periods (Unden et al., 1995). The easiest way to increase the amount of available oxygen in shake vessels is to increase shaking speed. For regular flasks, the optimal shaking speed range is 400–450 rpm. More agitation is generated in baffled flasks; under these conditions, 350–400 rpm are enough for good aeration. However, vigorous shaking can induce the formation of foam, which will lower oxygen transfer. For this reason, the addition of an antifoaming agent is recommended, although it was shown that antifoams can affect the growth rate of several microorganisms and the yield of recombinant protein (Routledge et al., 2011; Routledge, 2012). Also, proper aeration depends on the ratio of culture volume to vessel capacity. As a rule of thumb, the culture volume should be less or equal to 10% of the shaking flask capacity, although in our hands, protein production with culture volumes occupying 20% of the flask capacity was possible (Rosano et al., 2011). A strategy that can produce significant increases in cell density is fed-batch fermentation. This approach has a wide availability of tools and methods, but it is beyond the scope of this paper and is addressed elsewhere (Yamanè, 1984; Yee and Blanch, 1992; Moulton, 2013).

Two rarely discussed parameters in the process of recombinant protein production are the preparation of the starting culture and the time of induction. Most protocols call for diluting a saturated overnight preculture (dilution factor 1/100) into the larger culture (Sivashanmugam et al., 2009). However, leaky expression of the chosen system can lead to plasmid instability, which may result in a poor yield of target protein. Also, in the starter culture, cells can be in dissimilar metabolic states. Upon dilution into fresh media, cells will grow at different rates leading to irreproducible induction points (Huber et al., 2009). A proper preculture (cells in an active equalized growing phase) can be prepared by growing the overnight starter culture at 20–25°C or by using a slow-release system for glucose, among other methods (Busso et al., 2008; Huber et al., 2009; Sivashanmugam et al., 2009). After inoculation and further growth, the inducer is often added in mid-log phase because the culture is growing fast and protein translation is maximal. However, induction at early stationary phase is also possible (Ou et al., 2004). In fact, in some cases the target protein was more soluble when inducer was added at this stage (Galloway et al., 2003). Presumably, the reduced rate of protein synthesis may result in less aggregation in IBs, as we describe below.

Inclusion Bodies Formation

When a foreign gene is introduced in E. coli, spatio-temporal control of its expression is lost. The newly synthesized recombinant polypeptide is expressed in the microenvironment of E. coli, which may differ from that of the original source in terms of pH, osmolarity, redox potential, cofactors, and folding mechanisms. Also, in high level expression, hydrophobic stretches in the polypeptide are present at high concentrations and available for interaction with similar regions. All of these factors lead to protein instability and aggregation (Hartley and Kane, 1988; Carrio and Villaverde, 2002). These buildups of protein aggregates are known as IBs. IB formation results from an unbalanced equilibrium between protein aggregation and solubilization. So, it is possible to obtain a soluble recombinant protein by strategies that ameliorate the factors leading to IB formation (Carrio and Villaverde, 2001, 2002). One is to fuse the desired protein to a fusion partner that acts as a solubility enhancer. Some examples were already described in Section “Affinity Tags.” In some cases the generation of IB can be an advantage, especially if the protein can be refolded easily in vitro. If that is the case, conditions can be adjusted to favor the formation IB, providing a simple method for achieving a significant one-step purification of the expressed protein (Burgess, 2009; Basu et al., 2011).

Disulfide bond formation

For many recombinant proteins, the formation of correct disulfide bonds is vital for attaining their biologically active three-dimensional conformation. The formation of erroneous disulfide bonds can lead to protein misfolding and aggregation into IB. In E. coli, cysteine oxidation takes places in the periplasm, where disulfide bonds are formed in disulfide exchange reactions catalyzed by a myriad of enzymes, mainly from the Dsb family (Messens and Collet, 2006). By contrast, disulfide bond formation in the cytoplasm is rare, maybe because cysteine residues are part of catalytic sites in many enzymes. Disulfide bond formation at these sites may lead to protein inactivation, misfolding, and aggregation (Derman et al., 1993). The cytoplasm has a more negative redox potential and is maintained as a reducing environment by the thioredoxin–thioredoxin reductase (trxB) system and the glutaredoxin–glutaredoxin reductase (gor) system (Stewart et al., 1998). This situation has a huge impact in the production of recombinant proteins with disulfide bonds. One option would be to direct the protein to the periplasm, as we have discussed in Section “Protein Toxicity.”

Nevertheless, expression in the cytoplasm is still possible thanks to engineered E. coli strains that possess an oxidative cytoplasmic environment that favors disulfide bond formation (Derman et al., 1993). Worthy of mention are the Origami (Novagen) and SHuffle (NEB) strains. We described earlier the OrigamiTM strain, as having a trxB-gor- genotype in the K-12 background (as this double mutant is not viable, a suppressor mutation in the ahpC gene is necessary to maintain viability; Bessette et al., 1999). OrigamiTM is also available in the BL21(DE3) lacY (TunerTM, Novagen) background. Addition of the pRARE plasmid for the extra advantage of correcting codon bias resulted in the construction of the Rosetta-gamiTM B strain (Novagen). The SHuffle® T7 Express strain [BL21(DE3) background, NEB] goes a little bit further. Besides the trxB- and gor- mutations, it constitutively expresses a chromosomal copy of the disulfide bond isomerase DsbC (Lobstein et al., 2012). DsbC promotes the correction of mis-oxidized proteins into their correct form and is also a chaperone that can assist in the folding of proteins that do not require disulfide bonds. Due to the action of DsbC, less target protein aggregates into IB.

Chaperone co-expression/chemical chaperones and cofactor supplementation

Molecular chaperones lie at the heart of protein quality control, aiding nascent polypeptides to reach their final structure (Hartl and Hayer-Hartl, 2002). Other specialized types of chaperones, like ClpB, can disassemble unfolded polypeptides present in IB. The high level expression of recombinant proteins results in the molecular crowding of the cytosol and quality control mechanisms may be saturated in this situation (Carrio and Villaverde, 2002). One strategy for solving this problem is to stop protein expression by inducer removal after a centrifugation step and addition of fresh media supplemented with chloramphenicol, an inhibitor of protein synthesis. This allows recruitment of molecular chaperones to aid in the folding of newly synthesized recombinant polypeptides (Carrio and Villaverde, 2001; de Marco and De Marco, 2004).

Given their function, it is not surprising that efforts to inhibit IB formation were directed to the co-expression of individual or sets of molecular chaperones (Caspers et al., 1994; Nishihara et al., 2000; de Marco et al., 2007). Commercially, one of the most used systems is the chaperone plasmid set from Takara (Nishihara et al., 1998, 2000). This set consists of five plasmids (pACYC derivatives) which allow overexpression of different chaperones or combinations of them: (i) GroES-GroEL, (ii) DnaK/DnaJ/GrpE, (iii) (i) + (ii), (iv) trigger factor, (v) (i) + (iv). On the other hand, if such a system is not at hand, the natural network of chaperones can be induced by the addition of benzyl alcohol or heat shock, though the latter is not recommended (de Marco et al., 2005).

When proteins are purified from IB, urea-denatured and then refolded in vitro, addition of osmolytes (also called chemical chaperones) in the 0.1–1 M range of concentration increases the yield of soluble protein (Rudolph and Lilie, 1996; Clark, 1998; Tsumoto et al., 2003; Alibolandi and Mirzahoseini, 2011). This situation can be mimicked in vivo by supplementing the culture media with osmolytes such as proline, glycine-betaine, and trehalose (de Marco et al., 2005). Also, the folding pathways that lead to the correct final conformation and stabilization of the proper folded protein may require specific cofactors in the growth media, for example, metal ions (such as iron-sulfur and magnesium) and polypeptide cofactors. Addition of these compounds to the batch culture considerably increases the yield as well as the folding rate of soluble proteins (Sorensen and Mortensen, 2005).

Slowing down production rate

Slower rates of protein production give newly transcribed recombinant proteins time to fold properly. This was previously addressed when we discussed the role of translational pauses at rare codons and their impact in the production of recombinant proteins. Moreover, the reduction of cellular protein concentration favors proper folding. By far, the most commonly used way to lower protein synthesis is reducing incubation temperature (Schein and Noteborn, 1988; Vasina and Baneyx, 1997; Vera et al., 2007). Low temperatures decrease aggregation, which is favored at higher temperatures due to the temperature dependence of hydrophobic interactions (Baldwin, 1986; Makhatadze and Privalov, 1995; Schellman, 1997).

When IB formation is a problem, recombinant protein synthesis should be carried out in the range 15–25°C, though one report described successful expression at 4°C for 72 h (San-Miguel et al., 2013). However, when working at the lower end of the temperature range, slower growth and reduced synthesis rates can result in lower protein yields. Also, protein folding may be affected as the chaperone network may not be as efficient (McCarty and Walker, 1991; Mendoza et al., 2000; Strocchi et al., 2006). The ArticExpressTM (Stratagene) strain (B line) possesses the cold-adapted chaperonin Cpn60 and co-chaperonin Cpn10 from the psychrophilic bacterium Oleispira antarctica (Ferrer et al., 2004). The chaperonins display high refolding activities at temperatures of 4–12°C and confer an enhanced ability for E. coli to grow at lower temperatures (Ferrer et al., 2003).

Protein Inactivity

Obtaining a nice amount of soluble protein is not the end of the road. The protein may still be of bad quality; i.e., it does not have the activity it should. Incomplete folding could be the culprit in this scenario (Gonzalez-Montalban et al., 2007; Martinez-Alonso et al., 2008). In this case, the protein adopts a stable soluble conformation but the exact architecture of the active site is still unsuitable for activity. Some options already addressed can be helpful in these cases. Some proteins require small molecules or prosthetic groups to acquire their final folded conformation. Adding these compounds to the culture media can increase the yield and the quality of the expressed protein significantly (Weickert et al., 1999; Yang et al., 2003). Also, erroneous disulfide bond formation can lead to protein inactivity (Kurokawa et al., 2000). In addition, protein production at lower temperatures has a profound impact on protein quality. Work by the Villaverde lab has shown that conformational quality and functionality of highly soluble recombinant proteins increase when the temperature of the culture is reduced (Vera et al., 2007). This was also the case when the intracellular concentration of the chaperone DnaK was elevated (Martinez-Alonso et al., 2007). This phenomenon calls into question the use of solubility as an indicator of quality. Based on this fact, then it may be wise to express all recombinant proteins at low temperatures or at least, to compare the specific activity of a recombinant protein obtained at different temperatures.

If the activity of the heterologous protein is toxic to the cell, genetic reorganization of the expression vector leading to loss of activity may occur, allowing the host to survive and eventually take over the culture (Corchero and Villaverde, 1998). This structural instability of the plasmid can be detected by DNA sequencing after purification of the plasmid at the end of process. Any point mutation, deletion, insertion, or rearrangement may explain the low activity of a purified recombinant protein (Palomares et al., 2004).

Concluding Remarks

In terms of recombinant expression, E. coli has always been the preferred microbial cell factory. E. coli is a suitable host for expressing stably folded, globular proteins from prokaryotes and eukaryotes. Even though membrane proteins and proteins with molecular weights above 60 kDa are difficult to express, several reports have had success in this regard (our laboratory has produced proteins from plants in the 90–95 kDa range; Rosano et al., 2011). Large-scale protein expression trials have shown that <50% of bacterial proteins and <15% of non-bacterial proteins can be expressed in E. coli in a soluble form, which demonstrates the versatility of the system (Braun and LaBaer, 2003). However, when coming across a difficult-to-express protein, things can get complicated. We hope to have given a thorough list of possible solutions when facing the challenge of expressing a new protein in E. coli. Nevertheless, a word of caution is needed. Many of the approaches described in this review will fail miserably in a lot of cases. This can be explained by the fact that strategies aiming at troubleshooting recombinant protein expression are sometimes protein specific and suffer from positive bias; i.e., things that work get published, all the others, do not. That being said, thanks to the efforts of the scientific community, the general methods available in the literature are no longer anecdotal and can be used systematically. Moreover, the field is always expanding and even after almost 40 years from the first human protein obtained in E. coli (Itakura et al., 1977), there is still much room for improvement.

Author Contributions

Germán L. Rosano and Eduardo A. Ceccarelli wrote the manuscript and approved its final version.

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.


We would like to thank the reviewers for their insightful comments on the manuscript, as their remarks led to an improvement of the work. Germán L. Rosano and Eduardo A. Ceccarelli are staff members of the Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET, Argentina). Also, Germán L. Rosano is a Teaching Assistant and Eduardo A. Ceccarelli is a Professor of the Facultad de Ciencias Bioquímicas y Farmacéuticas, UNR, Argentina. This study was supported by grants from CONICET and Agencia Nacional de Promoción Científica y Tecnológica (ANPCyT, Argentina).


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Single Cell Protein—State-of-the-Art, Industrial Landscape and Patents 2001–2016


Humans and animals consume protein as a source of nitrogen and essential amino acids, from which they build new structural and functional (e.g., enzymes and hormones) proteins that enable them to survive. In extreme conditions, proteins may also be used as a source of energy. The nutritional value of a protein is determined by the amino acid composition; 20 amino acids are commonly found in dietary protein, of which several (i.e., phenylalanine, valine, threonine, tryptophan, methionine, leucine, isoleucine, lysine, and histidine, with arginine, cysteine, glycine, glutamine, proline, and tyrosine also being beneficial) cannot be synthesised by humans or animals and are thus essential and have to be supplied through the diet (for a review see Wu, 2009).

Boland et al. (2013) have explored how an increasing demand for meat and dairy protein will require improvements in animal production, as well as openness to new sources of protein, both as animal feed and for direct human consumption. Animal and dairy production have been increasing steadily over recent decades and theoretically can continue to do so to meet the expected demand, even by 2050 when the demand for meat would surpass 400 M tonnes and that for dairy 800 M tonnes (Boland et al., 2013). However, because plant protein is converted rather inefficiently into meat protein (~6 kg of plant protein is needed to produce 1 kg of meat protein), increasing meat production to match the growing demand is ultimately not sustainable (WHO, 2015). The western world is also interested in developing healthier food, with optimal amino acid composition and low, but good quality fat, combined with ethically sustainable production. These are typically non-animal based, environmentally friendly processes, but may include novel processes such as the production of “cultured” meat, in which meat protein is more efficiently produced in vitro rather than by growing an entire animal (Kadim et al., 2015). Plant-based protein sources, such as beans, are nutritionally valuable sources of protein, but require arable land and water, both of which will become limiting as we strive to meet global protein demand. The protein content of meat is generally about 45%, while that of milk is about 25% and of soybean about 35% (Ghasemi et al., 2011).

Protein can also be provided through the cultivation of various microbes and algae, preferably those which contain more than 30% protein in their biomass and which can provide a healthy balance of essential amino acids. Microbial protein is generally referred to as single cell protein (SCP), although some of the producing microbes, such as filamentous fungi or filamentous algae, may be multicellular. In addition to direct use as SCP, microbes contribute to protein demand when they are used to upgrade the protein content or quality of fermented foods (Bourdichon et al., 2012). Although, microbial protein provides a relatively small proportion of current human nutrition, the growing global demand for protein is likely to make SCP increasingly important (Boland et al., 2013). High growth rates or ability to utilise unique substrates, such as CO2 or methane, result in processes which offer much higher efficiency and/or sustainability than is possible from traditional agriculture.

SCP is currently produced from a limited number of microbial species, particularly when considering human consumption. The range of sources for SCP used in animal feed is broader than that approved for human consumption and is expanding. As is reviewed below, products derived from algae, fungi (including yeast) and bacteria are all in use or under development. The production steps generally include (a) preparation of nutrient media, possibly from waste, (b) cultivation, including solid state fermentation, (c) separation and concentration of SCP, in some cases drying, and (d) final processing of SCP into ingredients and products.

SCP for human consumption is generally produced from food grade substrates, but there is hope that processes will be developed to produce SCP from inexpensive waste materials from the food and beverage processing industries, as well as directly from forestry and agricultural sources (Anbuselvi et al., 2014). Regulatory issues must always be taken into account. With the introduction of algae to microbial protein providers, production from CO2 has become possible, while the greenhouse gas methane is providing a novel source of carbon for SCP from bacteria.

The following review will give an introduction to SCP production and the organisms used as SCP, with a focus on commercially implemented developments in the field. More detailed reviews of research with specific organisms considered for SCP production are provided by Anupama and Ravindra (2000), Ugalde and Castrillo (2002), Rudravaram et al. (2009), Ghasemi et al. (2011), and Nasseri et al. (2011). Here we provide an update on recent advances in the patent landscape (2001–2016) and the current industrial players, based on company profiles found from the web, literature and patent databases.

SCP Production Systems with Different Substrates and Processes

Algae, fungi (filamentous fungi and yeast), and bacteria can all be used as SCP (Anupama and Ravindra, 2000). In the future, dietary protein may also be derived from proteins secreted by engineered microbial cells (e.g., milk or egg white proteins) and produced from animal and plant cell cultures, in which the cells are no longer microbes but are not animals or plants, either. Thus, the distinction of what is SCP and what is other protein becomes blurred.

SCP from Algae

Microalgae which are produced for human or animal consumption typically have high protein content (e.g., 60–70%; Table 1). They also provide fats (with ω-3 fatty acids and carotenoids being of particular interest), vitamins A, B, C, and E, mineral salts, and chlorophyll (Gouveia et al., 2008). They have relatively low nucleic acid content (3–8%; Nasseri et al., 2011).

Table 1. Recent reports of the protein content of some algae that are of interest as SCP*.

Microalgae are currently used mainly in the form of supplements, available in tablet, capsule or liquid form, but they are increasingly also processed as ingredients which can be included in pastas, baked goods, snacks, and so on (Gouveia et al., 2008; Zimberoff, 2017). The most accessible commercial products are derived primarily from Arthrospira platensis and Arthrospira maxima (sold as spirulina, marketed by e.g., Hainan Simai Pharmacy Co., Earthrise Nutritionals, Cyanotech Corp., FEBICO, and Mayanmar Spriulina Factory), Chlorella (marketed by e.g., Taiwan Chlorella Manufacturing Co., FEBICO and Roquette Klötze GmbH & Co), Dunaliella salina (marketed by e.g., Qianqiu Biotechnology Co., Ltd., primarily for β-carotene) and Aphanizomenon flos-aquae (marketed by e.g., Blue Green Foods, Klamath Valley Botanicals LLC and E3Live; Gouveia et al., 2008). Euglena Co. Ltd. (Suzuki, 2017) and Algaeon ( are both selling products from Euglena, primarily for the β-glucan content, but including whole cell products. TerraVia does not specify the alga provided in their AlgaVia® food ingredient. Enzing et al. (2014) and Vigani et al. (2015) provide useful surveys of the companies and countries involved in production of microalgae as food or feed. Both reviews focus on the European Union, but take note of the involvement of numerous companies in Asia and North America in the industry.

Algae generally feed on CO2 and light, although some products such as AlgaVia® are produced by traditional fermentation rather than by photosynthesis. Outdoor production of algae in open ponds is common, but is subject to contamination (not only biological contamination, but also mineral contamination which affects the quality of the final product) and variation in the weather (Harun et al., 2010). Indoor photobioreactors are also being used to guarantee the supply of fresh algae as feed for aquaculture (molluscs, shrimp, fish; Henrikson, 2013; Mahmoud et al., 2016). Algae are primarily used in aquaculture as a source of omega fatty acids and carotenoid pigments, but their protein also contributes to animal nutrition (Muller-Feuga, 2000).

SCP from Fungi

A wide range of fungi have been considered for use as SCP, as reviewed by Anupama and Ravindra (2000), Rudravaram et al. (2009), and Nasseri et al. (2011). Table 2 lists some of the species that have been researched in recent years, with the protein content observed under the conditions in which they were grown. Products from Saccharomyces, Fusarium, and Torulopsis are commercially available.

Table 2. Recent reports of fungal protein content produced from specific substrates for species investigated as potential sources of SCP.

Fungi grown as SCP will generally contain 30–50% protein (Anupama and Ravindra, 2000; Nasseri et al., 2011). The amino acid composition compares favourably with the FAO guidelines; threonine and lysine content is typically high, but methionine content relatively low, although still meeting the FAO/WHO recommendations (Anderson et al., 1975). The methionine content of some fungal products such as Marmite® is even lower. Sulphur containing amino acids have been enriched in SCP from K. fragilis by cultivation on whey (Willetts and Ugalde, 1987).

In addition to protein, SCP derived from fungi is expected to provide vitamins primarily from the B-complex group (thiamine, riboflavin, biotin, niacin, pantothenic acid, pyridoxine, choline, streptogenin, glutathione, folic acid, and p-amino benzoic acid). The cell walls of fungi are rich in glucans, which contribute fibre to the diet. Low-density lipoprotein cholesterol has been reduced in volunteers who consumed myco-protein from Fusarium venenatum (Turnbull et al., 1992) and blood glucose and insulin levels may also be favourably affected (Lang et al., 1999). Fungi are expected to have a moderate nucleic acid content (7–10%; Nasseri et al., 2011), which however is too high for human consumption and requires processing to reduce it (Edelman et al., 1983).

The Quorn™ brand ( was launched in 1985 by Marlow Foods (UK). Quorn™ products contain mycoprotein from the filamentous fungus F. venenatum. The fungal biomass provides a texture that resembles meat products. Quorn™ may be the only SCP product exclusively used for human nutrition and has been extensively branded, marketed and sold for that purpose. The company was recently (2015) acquired by the Philippine instant noodles maker Monde Nissin Corp for 550 million pounds (

Spent brewer's yeast (Saccharomyces cerevisiae) have been sold for more than a century in yeast extracts such as Marmite® (Unilever and Sanitarium Health Food), Vegemite® (Bega Cheese Ltd.), Cenovis® (Gustav Gerig AG), and Vitam-R® (VITAM Hefe-Produkt GmbH). Yeast extracts provide a good source of five important group-B vitamins, but also protein. Another commercially available yeast, Torula (Candida utilis, renamed as Pichia jadinii), a widely used flavoring agent, is also high in protein. Torula was used in Provesteen® T, produced by the Provesta Corporation in the 1980s, along with similar products using Pichia and Kluyveromyces yeast (Hitzman, 1986). Torula is rich in the amino acid glutamic acid and for this reason it has been used to replace the flavor enhancer monosodium glutamate (MSG).

A process called “Pekilo” was developed in Finland to produce SCP for animal feed from the sugars present in sulphite liquor of paper mill effluents (reviewed in Ugalde and Castrillo, 2002). The filamentous fungus Paecilomyces varioti was grown on sugars, including pentoses, in the sulphite waste liquor or wood hydrolysates. There were two factories operating in Finland in Mänttä and in Jämsänkoski during 1982–1991, but as the cellulose mills ceased operations, these factories were also closed. Although, the product was sold as animal feed, it was also investigated as a supplement in meat products such as sausages and meat balls (Koivurinta et al., 1979). The Pekilo process strain is available from the VTT Ltd. culture collection (

Quorn™ and yeast spreads like Marmite® are produced from starch-derived glucose, while the Pekilo process used lignocellulosic sugars. In addition to these carbon sources, alkanes and methanol have been used for SCP production by yeast and filamentous fungi. Methylotrophic yeasts, for example Komagataella pastoris (previously Pichia pastoris), produce biomass and protein from methanol (Rashad et al., 1990). Industrial scale production has been carried out, e.g., by Phillips Petroleum Company. Their yeast produced 130 g (DW)/l biomass, with a productivity of more than 10 g l−1 h−1 (Johnson, 2013).

British Petroleum pioneered production of Yarrowia lipolytica SCP for animal feed from waxy n-paraffins from an oil refinery in the 1970's, building a pilot plant with up to 100 kton annual production capacity (Groenewald et al., 2014). Although the product itself was considered safe, the plant failed to get the required production permits because of environmental concerns (Bamberg, 2000). Combined with the high price of substrate resulting from the 1973 oil crises, this led British Petroleum to abandon its interest in SCP (Groenewald et al., 2014). Yarrowia SCP is now available on a smaller scale as Yarrowia Technology products (Yarrowia Equinox and Yarrowia GoodStart products) from Skotan S.A. in Poland ( Although, oils and carotenoids are the most common Yarrowia products for human use (Groenewald et al., 2014), the American-based Nucelis also offers a protein rich Yarrowia Flour (

Research and development on SCP with various fungal species is active and ongoing and may lead to novel products or production processes. For example, Zhao et al. (2013) described a process in which antibacterial peptides would be produced and secreted by Y. lipolytica, generating a high-value product, while the spent yeast could be used as SCP, since its protein content was high. Much of the current research focuses on the use of waste substrates such as sugarcane bagasse (e.g., Penicillium janthinellum with 46% protein, Rao et al., 2010), brewery's spent grains, hemicellulosic hydrolysate (e.g., Debaryomyces hansenii, White et al., 2008; Kluyveromyces marxianus, Aggelopoulos et al., 2014), whey (mixed yeast cultures, Yadav et al., 2014, 2016; K. marxianus, Aggelopoulos et al., 2014), and mixtures of other common food industry wastes such as orange and potato residues, molasses, and malt spent rootlets (K. marxianus, Aggelopoulos et al., 2014). Aggelopoulos et al. (2014) used solid state fermentation (SSF) rather than submerged cultivation and also noted that higher value products could be extracted prior to use of the protein-enriched residues as animal feed.

SCP from Bacteria

Bacteria also have a long history of use as SCP, particularly in animal feed. Some of the more commonly studied species have been reviewed by Anupama and Ravindra (2000), Rudravaram et al. (2009), and Nasseri et al. (2011) and Table 3 provides a list of more recent research on bacterial SCP.

Table 3. Recent reports of bacterial protein content on specific substrates for species investigated as potential sources of SCP.

Bacterial SCP generally contains 50–80% protein on a dry weight basis (Anupama and Ravindra, 2000) and the essential amino acid content is expected to be comparable to or higher than the FAO recommendations (Erdman et al., 1977). Methionine content up to 3.0% has been reported (Schulz and Oslage, 1976), which is higher than that generally obtained in algal or fungal SCP. Similar amino acid composition is observed with methanol or methane grown bacteria (Øverland et al., 2010). As with fungi, bacterial SCP has high nucleic acid content (8–12%), especially RNA, and thus requires processing prior to usage as food/feed (Kihlberg, 1972; Nasseri et al., 2011; Strong et al., 2015). In addition to protein and nucleic acid, bacterial SCP provides some lipid and vitamins from the B group.

Imperial Chemical Industries developed a SCP (Pruteen) for animal feed from methanol, using the bacterium Methylophilus methylotrophus. Pruteen contained up to 70% protein and was used in pig feed (Johnson, 2013). Pruteen, however, could not compete with cheaper animal feeds that were available at the end of the 1970s and production was discontinued. Pruteen was produced from methanol, but methane is now gaining interest as a substrate for SCP. UniBio A/S (utilizing knowledge gained by Dansk BioProtein A/S) and Calysta Inc. have both developed fermentation technology to convert natural gas to animal feed protein by using methanotrophic bacteria. UniBio A/S uses a U-loop fermenter, to achieve a productivity of 4 kg m−3 h−1, producing UniProtein® with ~70% protein, which has been approved for use in animal feed ( The U-loop fermenter is designed to enhance mass transfer rates of methane from the gas to the liquid phase, making more methane available for the bacteria (Petersen et al., 2017). Calysta Inc. opened a production facility for their product, FeedKind®, in the UK in 2016 and is partnering with Cargill to build a larger production facility in the U.S.A ( FeedKind®, like UniProtein®, is used in animal feed. Methane is an interesting substrate, since it is a major by-product of cattle and pig farming (Philippe and Nicks, 2015), as well as being available from biogas production (landfills, waste). Excess methane is currently flared. VTT Ltd. is investigating the reactor design and options for coupling farm methane generation with the production of microbial oil and feed protein ( from the methanotrophic bacteria Methylococcus capsulatus (group I), Methylosinus trichosporium (group II), and Methylocystis parvus (group II).

As with SCP from fungi, other developments in the production of bacterial SCP focus on upgrading various waste substrates or valorisation of waste water treatment. Examples include the treatment of potato starch processing waste in a two-step process using Aspergillus niger to degrade fibres in the potato residue and Bacillus licheniformis to produce protein (Liu et al., 2014). Economic analyses indicated that the process could address not only the pollution problem of the starch industry, but also the shortage of protein for animal feed in China (Liu et al., 2014). Another example of simultaneous waste water management and SCP production was reported by Kornochalert et al. (2014) for rubber sheet factory waste. They demonstrated that the chemical oxygen demand, suspended solids and total sulfides in the waste water was reduced by the purple nonsulfur bacterium, Rhodopseudomonas palustris, to levels that met the guidelines for use as irrigation water in Thailand and that the biomass produced was suitable for SCP (Kornochalert et al., 2014).

Soy-bean hull has been fermented with B. subtilis to improve its nutritional value as a feed for monogastric animals (Wongputtisin et al., 2014).

Kunasundari et al. (2013) describe a novel secondary product, co-produced with bacterial SCP. They cultivated Cupriavidus necator in a large scale to produce biomass high in both protein and polyhydroxyalkanoate (PHA). This biomass was fed to rats. The feed was not only well-tolerated and safe for rats, but the rats also produced faecal pellets containing PHA granules, which enabled the purification of substantial amounts of PHA without use of strong solvents (Kunasundari et al., 2013).

Processing of SCP

Depending on the substrate material and intended food/feed application, various processing steps are required prior to formulation of the final SCP product. In the following section we review the most relevant processing needs for SCP.

Cell Wall Degradation in Single Cell Protein Products

Some SCP are used as whole cell preparations, while in others the cell wall may be broken down to make the protein more accessible. SCP, such as Quorn™, may be consumed without degradation of the cell wall, in which case chitin and glucan from fungal cell walls contribute fibre to the diet (Wiebe, 2004). SCP derived from Euglena does not require dirsuption since the cells have proteinaceous pellicles, rather than cell walls, making it more readily digestible.

Various methods have been used to disrupt the cell wall, including mechanical forces (crushing, crumbling, grinding, pressure homogenization, or ultra-sonication), hydrolytic enzymes (endogenous or exogenous), chemical disruption with detergents, or combinations of these methods (reviewed in Nasseri et al., 2011). Cell disruption may affect the quality and quantity of protein and other components in the SCP. Products such as Marmite® and Vegemite® are cell extracts, generated by heating the cells to 45–50°C long enough for intracellular enzymes to partially hydrolyse the cell wall; the proteins are also reduced to smaller peptides (Trevelyan, 1976; Ugalde and Castrillo, 2002).

Nucleic Acid Removal in Single Cell Protein Products

Although algae generally have low nucleic acid content, the rapidly proliferating bacterial and fungal species have high nucleic acid (RNA) content. RNA content and degradation are affected by growth conditions, growth rate, and the carbon-nitrogen ratio (Trevelyan, 1976). When SCP is produced for human consumption, high nucleic acid content is a problem because ingestion of purine compounds derived from RNA breakdown increases uric acid concentrations in plasma, which can cause gout and kidney stones (Edelman et al., 1983). SCP with high nucleic acid content which is intended as animal feed is recommended only for feeding animals with short life spans (Strong et al., 2015). Gao and Xu (2015) and Xu (2015) have recently described methods for measuring the nucleotide content of complex SCP products.

Various methods to decrease the RNA content in SCP have been developed (Sinskey and Tannenbaum, 1975) and continue to be in use. Endogenous RNA degrading enzymes (ribonucleases) can be exploited in degradation of RNA, after activation with heat treatment (60–70°C) as used in the production of Quorn™ (Anderson and Solomons, 1984). Ribonucleases can also be added to the process or used as immobilized enzymes (Martinez et al., 1990; Hameş and Demir, 2015). Degraded RNA components diffuse out of the cells, but biomass loss (35–38%) also occurs. The process was improved by using higher temperatures (72–74°C) for 30–45 min, with less loss of biomass (30–33% loss; Ward, 1998). The temperature increase requires steam input, which is a cost factor, but heat is also needed for final treatment of the biomass at 90°C after the RNAse activation (Knight et al., 2001).

Alkaline hydrolysis and chemical extraction methods have also been studied. Viikari and Linko (1977) used an alkali treatment to reduce RNA in P. varioti biomass, used in for Pekilo-process, to below 2%. Treatment at 65°C, pH 7.5–8.5, to activate endogenous ribonuclease, also reduced the RNA content to <2%, while the protein content stayed at 50%.

Safety of SCPs

As for any food or feed product, SCP needs to be safe to produce and use. Regulations exist in most regions to ensure that food or feed are safe for consumption (Bagchi, 2006). Typically these distinguish not only between food (for humans) and feed (for animals), but also between food (providing nutrition and potentially taste and aroma) and food additives (preservatives, colourants, texture modifiers, etc.), or feed and feed additives. Exact definitions may differ between regions, but international standards, regulated through the Joint FAO/WHO Expert Committee on Food Additives, apply to internationally traded products (WHO, 2017). Regulations differ depending on the intended purpose of the product, and although SCP is expected to be either food or feed (providing nutrition), some products may enter the market as additives (e.g., providing colour), rather than as SCP, even though protein is present in the product, limiting the extent to which they are added and their value as SCP. Coppens et al. (2006) summarised the European regulations related to food and food supplements, concluding that “the process of having ‘functional foods’ ready for the market is certainly a costly and time-consuming task,” but also that the process can be successful.

Smedley (2013) provides useful references to the specific regulations related to feed and feed additives in Brazil, Canada, China, the European Union, Japan, South Africa, and the United States, and the differences between the regulations in these regions. It should be noted that not all animals are regarded the same in all regions, thus pet food is regulated as feed in some areas, but not in others. Authorisation is required before sale of new feed or additives (Smedley, 2013).

Key concerns are the RNA content, toxins produced by microbes (production hosts or contaminants), potential allergy symptoms, and harmful substances derived from the feedstock such as heavy metals. Methods have been developed and are in industrial use to decrease the RNA content to acceptable levels, as discussed above.

The challenge of toxins is overcome by carefully selecting the production organism, the process conditions, and the product formulation. Some fungi produce mycotoxins and this makes them undesirable sources of SCP (Anupama and Ravindra, 2000). The effects of fungal toxins range from allergic reactions to carcinogenesis and death. Both humans and animals are affected, so mycotoxins cannot be tolerated in SCP for either human or animal consumption. Quorn™ mycoprotein underwent extensive testing for the presence of mycotoxins or other toxic compounds before being approved for human consumption (Wiebe, 2004). The particular strain of F. venenatum does not produce mycotoxins under production conditions, but the process is still monitored to ensure none are present. The initial safety testing for Quorn™ mycoprotein involved 16 years, with many more years required to gain approval for sale outside the UK (Solomons, 1986). Y. lipolytica is another fungus whose safety has been extensively assessed, demonstrating that it would be safe to use in a variety of food applications, including as SCP (Groenewald et al., 2014).

Bacteria may also produce toxins which limit their use as SCP. Toxins may be extracellular (exotoxins) or cell bound (endotoxins). For example, both Pseudomonas spp. and Methylomonas methanica produce high levels of protein and have been assessed for use as SCP. Both also produce endotoxins that cause febrile reactions (Rudravaram et al., 2009). These can be destroyed by heating. Further, a study on immunogenicity of SCP from M. capsulatus showed that the cell-free preparation (i.e., the cell wall is removed) did not cause immune responses in mice, although whole cell preparations did (Steinmann et al., 1990).

The use of varying waste types of raw materials for SCP production is appealing from the cost and sustainability point of view, but may be challenging from the safety perspective and the origin of the feedstock must be carefully considered. For example, Quorn™ is produced in a chemically defined medium from glucose (hydrolysed starch) in a well-defined process which meets GLP standards (Wiebe, 2002, 2004). Any product for human consumption which would be produced from biomass hydrolysates or waste streams would need to provide an equivalent safety record before finding approval in Europe or North America. In addition to the safety requirements associated with the use of waste-derived substrates for SCP, public perception and acceptance of waste-derived foods would be a key element to consider when implementing SCPs in human diets.

Genetically Modified Organisms in SCP Production—Future Possibilities

Use of genetically modified organisms (GMO) in food and feed has not yet found public acceptance in Europe, although there is more acceptance elsewhere in the world. As data regarding GMO consumption accumulates, they may gain further acceptance as protein sources become scarcer, particularly if a market develops for healthy or personalized nutrition. GMO yeast from bioethanol factories can already be used as cattle feed in some countries. Use of genetic elements from the host itself (self-cloning) often means that no foreign DNA is introduced.

Although, Goldberg (1988) discussed the prospect of using genetically engineered microbes as SCP in the 1980s as a means of improving process economics by producing co-products (e.g., an enzyme, organic acid, or antibiotic), the concept was not pursued and has only gained more interest and acceptance in recent years. A wide range of advantages in SCP products from genetic modification has been considered. For example, DuPont has genetically engineered a yeast to produce long-chain omega-3 fatty acids, which are essential to human health (Xie et al., 2015). Genome sequencing and genetic engineering also allow disruption of genes involved in toxin production and thus improved safety of some SCP products. Disruption of genes can be achieved by traditional mutagenesis and screening, but the process may introduce undesired mutations into the product, whereas genetic modification is quicker and more specific. This will be aided by new technologies, such as Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) that allows specific editing of the genome without introduction of new DNA. Strains which have been modified using CRISPR are not necessarily considered GMOs. CRISPR methodology can also eliminate the introduction of antibiotic resistance genes to the organism, avoiding concern about the spread of antibiotic resistance genes through the use of GMOs.

Carbon source metabolism is another target for improving SCP production processes, since the carbon source can be a major cost in SCP manufacture (Ugalde and Castrillo, 2002). Genetic engineering could broaden the range of substrates used by the production organism or increase the efficiency of their use, enabling the use of multiple feed stocks and ensuring that all potential carbon in the raw material is used. For example, Ren et al. (2016) introduced xylose fermentation capability from Candida intermedia to S. cerevisiae by genome suffling to enable ethanol production from glucose and SCP production from xylose, while Cui et al. (2011) introduced inulase to Y. lipolytica. Similarly, expression of one or more hydrolytic enzymes has improved use of polymeric substrates (Song et al., 2017). Cellulose, starch, or whey could be used in consolidated bioprocesses by an organism modified to produce the tailor-made enzyme cocktail suitable for the particular raw material. Organisms could also be engineered to have improved tolerance to acid, alkali or other compounds associated with specific substrates.

Genetic modifications could also increase the nutraceutical value of the biomass, either by optimising the amino acid composition or by increasing the content of specific vitamins (e.g., D-vitamin, B-vitamin, biotin), fatty acids, glutathione, etc. along with the protein. There is considerable scope for creating SCPs with tailor made, personalized, nutritional composition.

Genetic engineering may also provide new ways of harvesting the proteins for inclusion in food or feed. For example, modification to improve flocculation could reduce costs in collection of cells or the cells could be modified to have a set of cell wall degrading enzymes that would be activated by specific extracellular stimuli to provide proteins without cell walls. Similarly, ribonucleases could be designed to be activated at a specific time in conditions in which proteases would not be activated. Morphological characteristics could also potentially be engineered to provide specific organoleptic properties.

Economic Aspects

Development of SCP processes has always been driven by a need for protein, and this continues to be an important driver in the development of both old and new processes. The valorisation of readily available substrate and waste streams has also been a strong driver and continues to be so. SCP is frequently seen as a potential co-product that could strengthen the economic potential of an otherwise unprofitable biorefinery process, as well as a means of reducing the downstream processing costs required to dispose of process waste. Selling residual biomass as feed is preferable to selling as fertiliser. This is seen in the numerous publications and patents (not addressed in this review) in which specific waste products are converted to SCP and are assessed as food for specific animals. However, environmental concerns also now play a strong role in driving the development of novel SCP products. This is seen particularly in the processes which utilise greenhouse gases: algal SCP from CO2 and bacterial SCP from methane. Such processes are unlikely to be economically viable in the short term, since there are still many problems to overcome in large scale cultivation, but may survive where they are able to benefit from a green premium. In addition, environmental concerns, as well as economic concerns, are helping to drive the development of products from waste streams.

Apart from the environmental benefits, the key elements in estimating the economic viability of a SCP production process are total product cost, capital investment and profitability. Ugalde and Castrillo (2002) estimated that in fungal SCP production 62% of the total product cost would come from the raw material and 19% from the production process. According to Aggelopoulos et al. (2014), raw material costs vary from 35 to 55% of the manufacturing costs, whereas the operation costs, including labour, energy, and consumables take 45–55%. Utilising side-streams and waste biomass is sometimes viewed as a means to reduce the substrate costs, in cases when the substrate does not compromise the usability of the final product.

Scale is also important to the economic viability of SCP production. An empirical relationship exists between cost and scale of production. Continuous operations have been proven to be the most profitable ones and the majority of the SCP processes which have been implemented at industrial scale have been adjusted to continuous design (Ugalde and Castrillo, 2002). On the other hand, small scale, household production of some products may become feasible, in much the way that home yoghurt production or mushroom production has, and as has been suggested for plant cell nutrition without plants (Poutanen et al., 2017).

Update on Industrial Production of SCP—Players and Capacities

Table 4, lists companies reported to produce or to have an interest in SCP, with website and patent information provided when available. A short description of some active companies is given below.

Algaeon Inc. produces β-glucan and whole cell products from the photosynthetic protist Euglena gracillis. Algaeon was started in 2011 and is based in the U.S.A.

BlueBioTech Int. GmbH, a microalgal biotechnology company, which has operated for more than 10 years, producing large quantities of Spirulina and Chlorella.

Calysta Inc. was founded as a private company in 2011. It produces FeedKind® from methane at a pilot facility in the UK, and began distributing commercial samples in 2017. It plans to open a larger facility (producing up to 20,000 tonnes per year) in the U.S.A. in 2019.

Cangzhou Tianyu Feed Additive Co., Ltd is a manufacturer and trading company located in Hebei, China since 2004. Their main products are Yeast Powder, Choline Chloride, Betaine, and Allicin having markets in Southeast Asia, Eastern Asia, Oceania, South Asia, and South America. The company employs 50 people and their total revenue is 5–10 million US$.

CBH Qingdao Co., Ltd has been an established company for decades supplying a range of ingredients and additives for feed and food industries. They can supply products which meet FAMI-QS, ISO, GMP, KOSHER, and HALAL standards.

Cyanotech Corporation is one of the world leading producers of Spirulina with sales in the US and 30 other countries. Their turnover in 2016 was almost 32 million US$. FDA has given GRAS status for Cyanotech's Spirulina as a food ingredient.

The progenitor of Earthrise, Proteus Corporation was founded in 1976. They produce Spirulina with GRAS status. They are GMP certified and have Food Safety System Certification (FSSC) 22000:2011.

E.I.D Parry Ltd., Parry Nutraceuticals Division is part of the 4.4 billion US$ Murugappa Group. They use micro-algal technology to produce nutraceuticals like Spirulina and Chlorella. Their products are sold in more than 40 countries and their main markets are in North America, Europe, South East Asia, and the Far East.

Euglena Co. Ltd. was founded in Japan in 2005. Amongst other products derived from Euglena gracillis, Euglena Co. Ltd. is developing de-fatted Euglena as a source of protein-rich animal feed.

KnipBio was founded in 2013 in the U.S.A. with a focus of providing affordable feed for aquaculture. They produce KnipBio Meal from methanol using a methylotrophic bacterium and plan to start commercial production in 2018.

Lallemand Inc. is a Canadian company specializing in the development, production, and marketing of yeast and bacteria. There are two major groups in the company: the Yeast Group (based in Montreal, Canada) and the Specialties Group (based in Toulouse, France). They produce SCP for human consumption (LBI, Lake States®, Engevita™) from the yeast S. cerevisiae and Torula.

LeSaffre produces yeast (S. cerevisiae) and yeast derived products including SCPs such as Lynside® Nutri, Lynside® ProteYn and related products (Lesaffre Human Care products), as well as yeast-based flavour ingredients (Biospringer products). The company has 7,700 employees and more than 80 subsidiaries in over 40 countries. Their products and services are sold in more than 180 countries and their turnover was ~1.6 billion € in 2013.

Marlow Foods Ltd produces the mycoprotein Quorn™. The Quorn development project started already in the 1960s, when they started to look for a microbial protein source that humans would find enjoyable. Quorn is classified as a safe, well-tolerated food by regulatory bodies across the world, including FDA, and the UK's Food Standards Agency (FSA). The company was acquired by Monde Nissin Corporation in the Philippines for 831 million US$ in 2015.

Nucelis Inc. was founded in 2010 in the U.S.A., but became a subsidiary of Cibus Global in 2014. Along with squalene, vitamin D and nutritional oils, Nucelis Inc. is developing high protein flour from the yeast Yarrowia.

Nutrinsic is based in the USA, with subsidiaries in China. Nutrinsic focuses on the use of waste waters from the food, beverage and biofuel industries to generate feed and fertiliser products. They market a SCP for animal feed called ProFloc™, which is described as having a protein content around 60%. They opened their first USA production facility in 2015, using waste water from a local brewery.

Tangshan Top Bio-Technology Co., Ltd is a manufacturer and trading company located in Hebei, China (Mainland). Their main products are: brewer's yeast, autolyzed yeast, yeast cell wall and yeast extract, including a 100% natural, non-GMO, pure yeast powder as animal feed additive for 1,100–1,250 US$ per ton and a production capacity of 15,000 tons per year per production line. The company was established in 2009 and has ~200 employees. Their main markets are in China, Eastern Asia, Western Europe, Southeast Asia and Mid East, with 40–50% of their products exported.

TerraVia Holdings, Inc. is a publicly held American company which focuses on providing ingredients for food and care products from eukaryotic algae. TerraVia appeared in 2016, but is derived from Solazyme Inc. which was founded in 2003. TerraVia uses traditional stirred tank reactors to cultivate its algae.

UniBio A/S, Denmark is an SME that owns rights to a unique fermentation technology—the U-Loop technology, which enables natural gas to be converted into a high protein product—UniProtein®. UniProtein® has a protein content of ~71% and can be used in feed for animals. UniBio A/S was established in 2001.

Unilever produces yeast extract Marmite® from brewer's spent grain. The number of employees is around 169,000 and the turnover of the company was $52.7 billion in 2016.

Vega Pharma Ltd is located in Zhejiang, China—the Vega Group is developing, manufacturing, and marketing pharmaceuticals, nutritional ingredients, animal health products, and probiotics. They offer a SCP, with up to 65% protein and containing relatively high threonine levels, for animal feed as a by-product of monosodium glutamate production.

Table 4. Industrial establishments involved in SCP production.

Recent Patents (2001–2016)

Recent patents (2001–2016) related to SCP production with algae, fungi, bacteria and mixed microbial populations are listed in Tables 5–8. Some of the patents owned by industrial operators are also shown in Table 4. The number of patents related to the use of algae, bacteria, yeast, or mixed populations is relatively evenly divided. Many patents have also been filed in which microbial biomass forms a component of a feed mixture which is intended to provide protein and other nutrients to fish or farmed animals. These have not been included in Tables 5–8, since it is not clear how much protein is provided by the microbe and how much by other components such as soy, bean, or fish meal.

Table 5. Patents related to the production of SCP from algae during 2001–2016*.

Table 6. Patents related to the production of SCP from yeast or filamentous fungi during 2001–2016*.

Table 7. Patents related to the production of SCP from bacteria during 2001–2016*.

Table 8. Patents related to the production of SCP from mixed microbial populations (bacteria and/or yeast and/or algae) or in which the microorganism was not specified during 2001–2016*.

Industries and universities in China have been particularly active in filing patents related to SCP in recent years, with about 70% of patents awarded since 2001 having been filed in China. In China, there has been a strong emphasis on the production of SCP by fermenting agricultural or food residues with bacteria, yeast and mixed populations. SCP production is thus often combined with bioremediation and waste processing.

Several important patents related to the use of C1 compounds such as methanol and methane were filed before 2001 and have not been included in this review. However, there were two new patents on the production of SCP from methanol and six on producing SCP from methane (Table 7). SCP from algae also continues to generate patents, with formulation of products attracting attention as well as continued developments in the cultivation methods (Table 5).

Concluding Remarks

As seen in Table 4, there are a wide range of industries involved in SCP production, some producing SCP as a by-product of other processes, and others which focus primarily on SCP. SCP from filamentous fungi and yeast continues to dominate the established markets, particularly when considering SCP for human consumption. Yeast SCPs have a long history of use, which facilitates their continued acceptance in the market. SCP for humans from filamentous fungi, however, is likely to remain restricted to F. venenatum (Quorn™) and solid-state fermentations with other food fungi, because of the risk of mycotoxins and the long path to regulatory acceptance. Yeast also have a long history of use as supplements to the feed industry. Much of the fungal SCP provided for animal feed is a byproduct of the food and beverage industries and of biorefineries, in which the fungus first acts as the biocatalyst to create the main product and then provides protein-enriched residues which are sold as feed. Fungal SCPs offer the advantages of familiarity, with well-established processing approaches, and availability. The main barrier is in the introduction of SCP from new species, which generate academic and patent interest, but which are difficult to bring into the market.

Algae also have well-established markets for both food and feed applications, although these are not traditionally focused on algae as SCP, but rather as food supplements providing omega-3 fatty acids, carotenoids and vitamins, with protein as a corollary benefit. Since products have been treated as supplements or colorants, the regulatory requirements are different than those for direct food or feed use, facilitating the introduction of new species for potential products. Algal products typically have flavours which may limit the amount a person would want to consume, reducing the need for extensive processing to reduce RNA, but also limiting the amount of protein provided to the diet. However, several of the young SMEs which have entered the market are developing processes to produce low-flavour products which could expand the contribution of algal protein in human diets. Algal SCP offers the advantages of providing healthy lipids along with the protein, while potentially consuming CO2. It has the benefit of being seen as environmentally friendly and very “green.” The main barriers are cost of production and the need for novel formulation to make it acceptable for humans. Algal SCP is likely to be strong in the feed industry, if the production costs can be reduced.

Bacterial SCP is primarily restricted to the feed industry, if not including cyanobacterial products with non-photosynthetic bacteria. Some bacterial SCP is currently a by-product of other industries such as monosodium glutamate production, and this type of feed product is expected to increase with the expansion of biorefineries, as with yeast. However, the most interesting current bacterial SCP developments relate to the use of methane as a carbon source. Although, the use of methane to produce bacterial biomass is not new, the drivers pushing developments have shifted from methane as a cheap carbon source to bacteria as a means of reducing green-house-gas emissions and the potential integration of feed production with animal farming. The low solubility of methane, coupled with low growth rates of the bacteria, poses a strong barrier to success in this area. However, young SMEs like UniBio and Calysta Inc. believe that the barriers can be overcome. Bacterial SCP, other than from methane, offers advantages in high production rates, but is disadvantaged by low familiarity and high nucleic acid content which adds to the processing costs.

SCP initially gained importance in human nutrition during times of war, when traditional sources of protein became scarce. It again became of interest during the latter half of the twentieth century because of concern about meeting the protein demands of the world's ever increasing population. These concerns were global, but when we consider current interest in SCP, we observe that the countries now driving research and development of new SCP are generally those with large populations (e.g., China and India) and problems with malnutrition. Most recent patents related to SCP have been filed from China, indicating the importance of SCP for future food and feed there. Fast growth of SCP products can be expected in China and perhaps in the whole of Asia. Development of algal SCP forms an exception to this observation—since many companies have been established around the world in recent years to develop products which can exploit the current excess availability of CO2. The drivers for development of algal SCP are thus somewhat different from those for the development of bacterial and fungal SCP. Production of SCP from methane shares this environmental concern and opportunity with the algal developments.

An increase in biorefinery processes, as part of the expansion of the bio-economy and circular economy concepts, also acts as a driver for the development of SCPs for use as animal feed, since conversion of waste material to animal feed offers better returns on investment than burning residual microbial biomass or utilising it as fertiliser. Regulatory clearance is still needed for use of novel products in animal feed, but this differs from that needed for human food use, and a wider range of substrates are considered acceptable when the product is intended for animal use. Thus, greater expansion in available SCPs for animal feed than for human food can be expected. None-the-less, there is a growing appreciation of the inefficiency of converting plant biomass into SCP which is fed to animals, rather than directly to humans, which will push development of safe SCP as food also.

In the west, interest in healthy diets and novelty food is helping to drive a new interest in SCP, while also blurring the edges of what products might be included in SCP. Cell cultures of both plant and animal cells may contribute to food supply in the future (Poutanen et al., 2017), but do not conform to the definition of SCP as being derived from microbial cells. In addition, the forms in which SCP may be consumed are continuing to evolve. Yeast SCP has been consumed for decades as a cell extract in the form of pastes which can be spread on bread, whereas the fungal SCP which is used in Quorn™ was deliberately developed as a product which could be formulated into chunks and slices which would more closely resemble meat. More recently developed products are often formulated as dry powders or flours, which are intended to be mixed with other ingredients to create products in which the individual components are not perceived. Such products are suitable for incorporation into protein bars and beverages such as smoothies, which are currently popular. Additionally, solid state fermentations continue to be developed which use microbes to upgrade the protein quality and palatability of low nutrient plant products or ingredients. These are not strictly speaking SCP, since both the microbe and the original substrate contribute to the final product, but they will also contribute to the protein supply of the future. Having a broad range of food products which incorporate SCP should encourage further expansion of the market.

Author Contributions

AR, SH, and MT contributed equally to the researching and writing of this article. MW provided information on algal and fungal SCP, contributed to writing the article and reviewed and edited the manuscript.


The authors thank VTT Ltd. for financial support.

Conflict of Interest Statement

The authors are employees of VTT Technical Research Centre of Finland Ltd. and declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. MW received financial support from Marlow Foods in 1986-1989 and worked in projects supported by Marlow Foods from 1989 to 1994, but has had no ongoing collaboration with them.

The other authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.


The authors thank Drs. Johanna Buchert, Laura Ruohonen, and Tiina Nakari-Setälä for requesting the review and encouraging its publication.


DSP, Downstream processing; GMO, Genetically modified organism; GRAS, Generally recognised as safe; SCP, Single cell protein; QPS, Qualified presumption of safety of micro-organisms in food and feed applications.


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Keywords: single cell protein, SCP, algae, bacteria, fungi, microbial protein, Quorn™

Citation: Ritala A, Häkkinen ST, Toivari M and Wiebe MG (2017) Single Cell Protein—State-of-the-Art, Industrial Landscape and Patents 2001–2016. Front. Microbiol. 8:2009. doi: 10.3389/fmicb.2017.02009

Received: 03 August 2017; Accepted: 29 September 2017;
Published: 13 October 2017.

Edited by:

Andrea Gomez-Zavaglia, Center for Research and Development in Food Cryotechnology (CIDCA, CONICET), Argentina

Copyright © 2017 Ritala, Häkkinen, Toivari and Wiebe. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

*Correspondence: Marilyn G. Wiebe, [email protected]

Protein Structure and Folding

Learning Objectives

  • Describe the fundamental structure of an amino acid
  • Describe the chemical structures of proteins
  • Summarize the unique characteristics of proteins

At the beginning of this chapter, a famous experiment was described in which scientists synthesized amino acids under conditions simulating those present on earth long before the evolution of life as we know it. These compounds are capable of bonding together in essentially any number, yielding molecules of essentially any size that possess a wide array of physical and chemical properties and perform numerous functions vital to all organisms. The molecules derived from amino acids can function as structural components of cells and subcellular entities, as sources of nutrients, as atom- and energy-storage reservoirs, and as functional species such as hormones, enzymes, receptors, and transport molecules.

Amino Acids and Peptide Bonds

A table titled some amino acids and their structures; 3 columns: amino acid, R group, structure. Alanine has an R group of CH3. Its structure is a C attached to a COO-, an H, a NH3, and a CH3. Serine has an R group of CH2OH. Its structure is a C attached to a COO-, an H, a NH3, and a CH2OH. Lysine has an R group of (CH2)4NH3+. Its structure is a C attached to a COO-, an H, a NH3, and a (CH2)4NH3+. Aspartate has an R group of CH2COO. Its structure is a C attached to a COO-, an H, a NH3, and a CH2COO. Cysteine has an R group of CH2SH. Its structure is a C attached to a COO-, an H, a NH3, and a CH2SH.

Figure 1.

An amino acid is an organic molecule in which a hydrogen atom, a carboxyl group (–COOH), and an amino group (–NH2) are all bonded to the same carbon atom, the so-called α carbon. The fourth group bonded to the α carbon varies among the different amino acids and is called a residue or a side chain, represented in structural formulas by the letter R. A residue is a monomer that results when two or more amino acids combine and remove water molecules. The primary structure of a protein, a peptide chain, is made of amino acid residues. The unique characteristics of the functional groups and R groups allow these components of the amino acids to form hydrogen, ionic, and disulfide bonds, along with polar/nonpolar interactions needed to form secondary, tertiary, and quaternary protein structures. These groups are composed primarily of carbon, hydrogen, oxygen, nitrogen, and sulfur, in the form of hydrocarbons, acids, amides, alcohols, and amines. A few examples illustrating these possibilities are provided in Figure 1.

Amino acids may chemically bond together by reaction of the carboxylic acid group of one molecule with the amine group of another. This reaction forms a peptide bond and a water molecule and is another example of dehydration synthesis (Figure 2). Molecules formed by chemically linking relatively modest numbers of amino acids (approximately 50 or fewer) are called peptides, and prefixes are often used to specify these numbers: dipeptides (two amino acids), tripeptides (three amino acids), and so forth. More generally, the approximate number of amino acids is designated: oligopeptides are formed by joining up to approximately 20 amino acids, whereas polypeptides are synthesized from up to approximately 50 amino acids. When the number of amino acids linked together becomes very large, or when multiple polypeptides are used as building subunits, the macromolecules that result are called proteins. The continuously variable length (the number of monomers) of these biopolymers, along with the variety of possible R groups on each amino acid, allows for a nearly unlimited diversity in the types of proteins that may be formed.

Alanine has a 3 carbon chain. The second carbon has NH2 attached and the third has a double bonded O. When 2 alanines bond, the OH from one and the H from the NH2 of the other form water. The resulting molecule is two alanines linked by an NH.

Figure 2. Peptide bond formation is a dehydration synthesis reaction. The carboxyl group of the first amino acid (alanine) is linked to the amino group of the incoming second amino acid (alanine). In the process, a molecule of water is released.

Think about It

  • How many amino acids are in polypeptides?

Protein Structure

The size (length) and specific amino acid sequence of a protein are major determinants of its shape, and the shape of a protein is critical to its function. For example, in the process of biological nitrogen fixation (see Biogeochemical Cycles), soil microorganisms collectively known as rhizobia symbiotically interact with roots of legume plants such as soybeans, peanuts, or beans to form a novel structure called a nodule on the plant roots. The plant then produces a carrier protein called leghemoglobin, a protein that carries nitrogen or oxygen. Leghemoglobin binds with a very high affinity to its substrate oxygen at a specific region of the protein where the shape and amino acid sequence are appropriate (the active site). If the shape or chemical environment of the active site is altered, even slightly, the substrate may not be able to bind as strongly, or it may not bind at all. Thus, for the protein to be fully active, it must have the appropriate shape for its function.

Protein structure is categorized in terms of four levels: primary, secondary, tertiary, and quaternary. The primary structure is simply the sequence of amino acids that make up the polypeptide chain. Figure 3 depicts the primary structure of a protein.

The primary protein structure is a chain of amino acids that makes up the protein. The image is a chain of circles (each circle is an amino acid). One end of the chain is the free amino group or N-terminus. The other end of the chain is the free carboxyl group or C-terminus. A drawing of a single amino acid shows a carbon with an H, an R group, a COOH (acidic carboxyl group) and an NH2 (amino group).

Figure 3. Click to view a larger image. The primary structure of a protein is the sequence of amino acids. (credit: modification of work by National Human Genome Research Institute)

The chain of amino acids that defines a protein’s primary structure is not rigid, but instead is flexible because of the nature of the bonds that hold the amino acids together. When the chain is sufficiently long, hydrogen bonding may occur between amine and carbonyl functional groups within the peptide backbone (excluding the R side group), resulting in localized folding of the polypeptide chain into helices and sheets. These shapes constitute a protein’s secondary structure. The most common secondary structures are the α-helix and β-pleated sheet. In the α-helix structure, the helix is held by hydrogen bonds between the oxygen atom in a carbonyl group of one amino acid and the hydrogen atom of the amino group that is just four amino acid units farther along the chain. In the β-pleated sheet, the pleats are formed by similar hydrogen bonds between continuous sequences of carbonyl and amino groups that are further separated on the backbone of the polypeptide chain (Figure 4).

The secondary structure of a protein may be an α-helix or a β-pleated sheet, or both. A chain of spheres forms a spiral labeled alpha-helix. This chain also forms a ribbon that folds back and forth; this is labeled beta-pleated sheet. Closeups show that hydrogen bonds (dotted lines) between amino acids hold together these shapes.

Figure 4. The secondary structure of a protein may be an α-helix or a β-pleated sheet, or both.

A long ribbon labeled polypeptide backbone. Loops of the ribbon are held in place by various types of chemical reactions. An ionic bond is then a positively charged amino acid and a negatively charged amino acid are attracted to each other. Hydrophobic interactions are when hydrophobic amino acids (containing only carbons and hydrogens) are clustered together. A disulfide linkage is when a sulfur of one amino acid is covalently bound to the sulfur of another amino acid. A hydrogen bond is when two polar amino acids are attracted to each other.

Figure 5. Click to view larger image. The tertiary structure of proteins is determined by a variety of attractive forces, including hydrophobic interactions, ionic bonding, hydrogen bonding, and disulfide linkages.

The next level of protein organization is the tertiary structure, which is the large-scale three-dimensional shape of a single polypeptide chain. Tertiary structure is determined by interactions between amino acid residues that are far apart in the chain. A variety of interactions give rise to protein tertiary structure, such as disulfide bridges, which are bonds between the sulfhydryl (–SH) functional groups on amino acid side groups; hydrogen bonds; ionic bonds; and hydrophobic interactions between nonpolar side chains. All these interactions, weak and strong, combine to determine the final three-dimensional shape of the protein and its function (Figure 5).

The process by which a polypeptide chain assumes a large-scale, three-dimensional shape is called protein folding. Folded proteins that are fully functional in their normal biological role are said to possess a native structure. When a protein loses its three-dimensional shape, it may no longer be functional. These unfolded proteins are denatured. Denaturation implies the loss of the secondary structure and tertiary structure (and, if present, the quaternary structure) without the loss of the primary structure.

Some proteins are assemblies of several separate polypeptides, also known as protein subunits. These proteins function adequately only when all subunits are present and appropriately configured. The interactions that hold these subunits together constitute the quaternary structure of the protein. The overall quaternary structure is stabilized by relatively weak interactions. Hemoglobin, for example, has a quaternary structure of four globular protein subunits: two α and two β polypeptides, each one containing an iron-based heme (Figure 6).

A complex spherical shape made of ribbons that are coiled and wound around each other. There are 4 large regions (each made from a separate ribbon) – alpha 1, alpha 2, beta 1, beta 2. There are also red spheres attached to each ribbon; these are labeled heme group.

Figure 6. A hemoglobin molecule has two α and two β polypeptides together with four heme groups.

Another important class of proteins is the conjugated proteins that have a nonprotein portion. If the conjugated protein has a carbohydrate attached, it is called a glycoprotein. If it has a lipid attached, it is called a lipoprotein. These proteins are important components of membranes. Figure 7 summarizes the four levels of protein structure.

Primary protein structure: sequence of a chain of amino acids. This is shown as a chain of circles. Secondary protein structure: local folding of the polypeptide chain into helices or sheets. This is shown as a spiral labeled alpha-helix and a folded sheet labeled beta-pleated sheet. Tertiary protein structure: three-dimensional folding pattern of a protein due to side chain interactions. This is shown as a complex 3-D shape made of alpha helices and beta pleated sheets. Quaternary protein structure: protein consisting of more than one amino acid chain. This is shown as 2 complex structures similar to that seen at the tertiary level.

Figure 7. Protein structure has four levels of organization. (credit: modification of work by National Human Genome Research Institute)

Think about It

  • What can happen if a protein’s primary, secondary, tertiary, or quaternary structure is changed?

Primary Structure, Dysfunctional Proteins, and Cystic Fibrosis

A drawing of a phospholipid bilayer in the center with two protein channels. One is open and lets Cl- flow out of the cell. The other is blocked by a mucus blockage on the outside of the cell; Cl- ions can’t flow through this channel.

Figure 8. Click to view a larger image. The normal CFTR protein is a channel protein that helps salt (sodium chloride) move in and out of cells.

Proteins associated with biological membranes are classified as extrinsic or intrinsic. Extrinsic proteins, also called peripheral proteins, are loosely associated with one side of the membrane. Intrinsic proteins, or integral proteins, are embedded in the membrane and often function as part of transport systems as transmembrane proteins. Cystic fibrosis (CF) is a human genetic disorder caused by a change in the transmembrane protein. It affects mostly the lungs but may also affect the pancreas, liver, kidneys, and intestine. CF is caused by a loss of the amino acid phenylalanine in a cystic fibrosis transmembrane protein (CFTR). The loss of one amino acid changes the primary structure of a protein that normally helps transport salt and water in and out of cells (Figure 8).

The change in the primary structure prevents the protein from functioning properly, which causes the body to produce unusually thick mucus that clogs the lungs and leads to the accumulation of sticky mucus. The mucus obstructs the pancreas and stops natural enzymes from helping the body break down food and absorb vital nutrients.

In the lungs of individuals with cystic fibrosis, the altered mucus provides an environment where bacteria can thrive. This colonization leads to the formation of biofilms in the small airways of the lungs. The most common pathogens found in the lungs of patients with cystic fibrosis are Pseudomonas aeruginosa (Figure 9a) and Burkholderia cepacia. Pseudomonas differentiates within the biofilm in the lung and forms large colonies, called “mucoid” Pseudomonas. The colonies have a unique pigmentation that shows up in laboratory tests (Figure 9b) and provides physicians with the first clue that the patient has CF (such colonies are rare in healthy individuals).

a) a micrograph of rod shaped cells. B) An agar plate with a green pigmented colonies; this green pigment is spreading past the edge of the colonies.

Figure 9. (a) A scanning electron micrograph shows the opportunistic bacterium Pseudomonas aeruginosa. (b) Pigment-producing P. aeruginosa on cetrimide agar shows the green pigment called pyocyanin. (credit a: modification of work by the Centers for Disease Control and Prevention)

For more information about cystic fibrosis, visit the Cystic Fibrosis Foundation website.

Key Concepts and Summary

  • Amino acids are small molecules essential to all life. Each has an α carbon to which a hydrogen atom, carboxyl group, and amine group are bonded. The fourth bonded group, represented by R, varies in chemical composition, size, polarity, and charge among different amino acids, providing variation in properties.
  • Peptides are polymers formed by the linkage of amino acids via dehydration synthesis. The bonds between the linked amino acids are called peptide bonds. The number of amino acids linked together may vary from a few to many.
  • Proteins are polymers formed by the linkage of a very large number of amino acids. They perform many important functions in a cell, serving as nutrients and enzymes; storage molecules for carbon, nitrogen, and energy; and structural components.
  • The structure of a protein is a critical determinant of its function and is described by a graduated classification: primary, secondary, tertiary, and quaternary. The native structure of a protein may be disrupted by denaturation, resulting in loss of its higher-order structure and its biological function.
  • Some proteins are formed by several separate protein subunits, the interaction of these subunits composing the quaternary structure of the protein complex.
  • Conjugated proteins have a nonpolypeptide portion that can be a carbohydrate (forming a glycoprotein) or a lipid fraction (forming a lipoprotein). These proteins are important components of membranes.

Multiple Choice

Which of the following groups varies among different amino acids?

  1. hydrogen atom
  2. carboxyl group
  3. R group
  4. amino group
Show Answer

Answer c. The R group varies among different amino acids.

The amino acids present in proteins differ in which of the following?

  1. size
  2. shape
  3. side groups
  4. all of the above
Show Answer

Answer d. The amino acids present in proteins differ in all of the options: size, shape, and side groups.

Which of the following bonds are not involved in tertiary structure?

  1. peptide bonds
  2. ionic bonds
  3. hydrophobic interactions
  4. hydrogen bonds
Show Answer

Answer a. Peptide bonds are not involved in tertiary structure.

Fill in the Blank

The sequence of amino acids in a protein is called its __________.

Show Answer

The sequence of amino acids in a protein is called its Primary structure.

Denaturation implies the loss of the __________ and __________ structures without the loss of the __________ structure.

Show Answer

Denaturation implies the loss of the secondary and tertiary structures without the loss of the primary structure.


A change in one amino acid in a protein sequence always results in a loss of function.

Think about It

  1. Heating a protein sufficiently may cause it to denature. Considering the definition of denaturation, what does this statement say about the strengths of peptide bonds in comparison to hydrogen bonds?
  2. The image shown represents a tetrapeptide.A green 5-C chain linked to an NH linked to a black 2 C chain linked to an NH linked to a black 2 C chain linked to an NH linked to a blue 5 C chain.
    1. How many peptide bonds are in this molecule?
    2. Identify the side groups of the four amino acids composing this peptide.

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    Chemistry, nutrition, and microbiology of D-amino acids

    Exposure of food proteins to certain processing conditions induces two major chemical changes: racemization of all L-amino acids to D-isomers and concurrent formation of cross-linked amino acids such as lysinoalanine. Racemization of L-amino acids residues to their D-isomers in food and other proteins is pH-, time-, and temperature-dependent. Although racemization rates of the 18 different L-amino acid residues in a protein vary, the relative rates in different proteins are similar. The diet contains both processing-induced and naturally formed D-amino acids. The latter include those found in microorganisms, plants, and marine invertebrates. Racemization impairs digestibility and nutritional quality. The nutritional utilization of different D-amino acids varies widely in animals and humans. In addition, some D-amino acids may be both beneficial and deleterious. Thus, although D-phenylalanine in an all-amino-acid diet is utilized as a nutritional source of L-phenylalanine, high concentrations of D-tyrosine in such diets inhibit the growth of mice. Both D-serine and lysinoalanine induce histological changes in the rat kidney. The wide variation in the utilization of D-amino acids is illustrated by the fact that whereas D-methionine is largely utilized as a nutritional source of the L-isomer, D-lysine is totally devoid of any nutritional value. Similarly, although L-cysteine has a sparing effect on L-methionine when fed to mice, D-cysteine does not. Because D-amino acids are consumed by animals and humans as part of their normal diets, a need exists to develop a better understanding of their roles in nutrition, food safety, microbiology, physiology, and medicine. To contribute to this effort, this multidiscipline-oriented overview surveys our present knowledge of the chemistry, nutrition, safety, microbiology, and pharmacology of D-amino acids. Also covered are the origin and distribution of D-amino acids in the food chain and in body fluids and tissues and recommendations for future research in each of these areas. Understanding of the integrated, beneficial effects of D-amino acids against cancer, schizophrenia, and infection, and overlapping aspects of the formation, occurrence, and biological functions of D-amino should lead to better foods and improved human health.


    Now discussing:


    Biological molecule consisting of chains of amino acid residues

    This article is about a class of molecules. For protein as a nutrient, see Protein (nutrient). For other uses, see Protein (disambiguation).

    Proteins are large biomolecules and macromolecules that comprise one or more long chains of amino acidresidues. Proteins perform a vast array of functions within organisms, including catalysing metabolic reactions, DNA replication, responding to stimuli, providing structure to cells and organisms, and transporting molecules from one location to another. Proteins differ from one another primarily in their sequence of amino acids, which is dictated by the nucleotide sequence of their genes, and which usually results in protein folding into a specific 3D structure that determines its activity.

    A linear chain of amino acid residues is called a polypeptide. A protein contains at least one long polypeptide. Short polypeptides, containing less than 20–30 residues, are rarely considered to be proteins and are commonly called peptides, or sometimes oligopeptides. The individual amino acid residues are bonded together by peptide bonds and adjacent amino acid residues. The sequence of amino acid residues in a protein is defined by the sequence of a gene, which is encoded in the genetic code. In general, the genetic code specifies 20 standard amino acids; but in certain organisms the genetic code can include selenocysteine and—in certain archaea—pyrrolysine. Shortly after or even during synthesis, the residues in a protein are often chemically modified by post-translational modification, which alters the physical and chemical properties, folding, stability, activity, and ultimately, the function of the proteins. Some proteins have non-peptide groups attached, which can be called prosthetic groups or cofactors. Proteins can also work together to achieve a particular function, and they often associate to form stable protein complexes.

    Once formed, proteins only exist for a certain period and are then degraded and recycled by the cell's machinery through the process of protein turnover. A protein's lifespan is measured in terms of its half-life and covers a wide range. They can exist for minutes or years with an average lifespan of 1–2 days in mammalian cells. Abnormal or misfolded proteins are degraded more rapidly either due to being targeted for destruction or due to being unstable.

    Like other biological macromolecules such as polysaccharides and nucleic acids, proteins are essential parts of organisms and participate in virtually every process within cells. Many proteins are enzymes that catalyse biochemical reactions and are vital to metabolism. Proteins also have structural or mechanical functions, such as actin and myosin in muscle and the proteins in the cytoskeleton, which form a system of scaffolding that maintains cell shape. Other proteins are important in cell signaling, immune responses, cell adhesion, and the cell cycle. In animals, proteins are needed in the diet to provide the essential amino acids that cannot be synthesized. Digestion breaks the proteins down for use in the metabolism.

    Proteins may be purified from other cellular components using a variety of techniques such as ultracentrifugation, precipitation, electrophoresis, and chromatography; the advent of genetic engineering has made possible a number of methods to facilitate purification. Methods commonly used to study protein structure and function include immunohistochemistry, site-directed mutagenesis, X-ray crystallography, nuclear magnetic resonance and mass spectrometry.

    History and etymology

    Further information: History of molecular biology

    Proteins were recognized as a distinct class of biological molecules in the eighteenth century by Antoine Fourcroy and others, distinguished by the molecules' ability to coagulate or flocculate under treatments with heat or acid.[1] Noted examples at the time included albumin from egg whites, blood serum albumin, fibrin, and wheat gluten.

    Proteins were first described by the Dutch chemist Gerardus Johannes Mulder and named by the Swedish chemist Jöns Jacob Berzelius in 1838.[2][3] Mulder carried out elemental analysis of common proteins and found that nearly all proteins had the same empirical formula, C400H620N100O120P1S1.[4] He came to the erroneous conclusion that they might be composed of a single type of (very large) molecule. The term "protein" to describe these molecules was proposed by Mulder's associate Berzelius; protein is derived from the Greek word πρώτειος (proteios), meaning "primary",[5] "in the lead", or "standing in front",[6] + -in. Mulder went on to identify the products of protein degradation such as the amino acidleucine for which he found a (nearly correct) molecular weight of 131 Da.[4] Prior to "protein", other names were used, like "albumins" or "albuminous materials" (Eiweisskörper, in German).[7]

    Early nutritional scientists such as the German Carl von Voit believed that protein was the most important nutrient for maintaining the structure of the body, because it was generally believed that "flesh makes flesh."[8]Karl Heinrich Ritthausen extended known protein forms with the identification of glutamic acid. At the Connecticut Agricultural Experiment Station a detailed review of the vegetable proteins was compiled by Thomas Burr Osborne. Working with Lafayette Mendel and applying Liebig's law of the minimum in feeding laboratory rats, the nutritionally essential amino acids were established. The work was continued and communicated by William Cumming Rose. The understanding of proteins as polypeptides came through the work of Franz Hofmeister and Hermann Emil Fischer in 1902.[9][10] The central role of proteins as enzymes in living organisms was not fully appreciated until 1926, when James B. Sumner showed that the enzyme urease was in fact a protein.[11]

    The difficulty in purifying proteins in large quantities made them very difficult for early protein biochemists to study. Hence, early studies focused on proteins that could be purified in large quantities, e.g., those of blood, egg white, various toxins, and digestive/metabolic enzymes obtained from slaughterhouses. In the 1950s, the Armour Hot Dog Co. purified 1 kg of pure bovine pancreatic ribonuclease A and made it freely available to scientists; this gesture helped ribonuclease A become a major target for biochemical study for the following decades.[4]

    Linus Pauling is credited with the successful prediction of regular protein secondary structures based on hydrogen bonding, an idea first put forth by William Astbury in 1933.[12] Later work by Walter Kauzmann on denaturation,[13][14] based partly on previous studies by Kaj Linderstrøm-Lang,[15] contributed an understanding of protein folding and structure mediated by hydrophobic interactions.

    The first protein to be sequenced was insulin, by Frederick Sanger, in 1949. Sanger correctly determined the amino acid sequence of insulin, thus conclusively demonstrating that proteins consisted of linear polymers of amino acids rather than branched chains, colloids, or cyclols.[16] He won the Nobel Prize for this achievement in 1958.[17]

    The first protein structures to be solved were hemoglobin and myoglobin, by Max Perutz and Sir John Cowdery Kendrew, respectively, in 1958.[18][19] As of 2017[update], the Protein Data Bank has over 126,060 atomic-resolution structures of proteins.[20] In more recent times, cryo-electron microscopy of large macromolecular assemblies[21] and computational protein structure prediction of small protein domains[22] are two methods approaching atomic resolution.

    Number of proteins encoded in genomes

    The number of proteins encoded in a genome roughly corresponds to the number of genes (although there may be a significant number of genes that encode RNA of protein, e.g. ribosomal RNAs). Viruses typically encode a few to a few hundred proteins, archaea and bacteria a few hundred to a few thousand, while eukaryotes typically encode a few thousand up to tens of thousands of proteins (see genome size for a list of examples).


    Chemical structure of the peptide bond (bottom) and the three-dimensional structure of a peptide bond between an alanineand an adjacent amino acid (top/inset). The bond itself is made of the CHONelements.

    Main articles: Biochemistry, Amino acid, and Peptide bond

    Most proteins consist of linear polymers built from series of up to 20 different L-α- amino acids. All proteinogenic amino acids possess common structural features, including an α-carbon to which an amino group, a carboxyl group, and a variable side chain are bonded. Only proline differs from this basic structure as it contains an unusual ring to the N-end amine group, which forces the CO–NH amide moiety into a fixed conformation.[23] The side chains of the standard amino acids, detailed in the list of standard amino acids, have a great variety of chemical structures and properties; it is the combined effect of all of the amino acid side chains in a protein that ultimately determines its three-dimensional structure and its chemical reactivity.[24] The amino acids in a polypeptide chain are linked by peptide bonds. Once linked in the protein chain, an individual amino acid is called a residue, and the linked series of carbon, nitrogen, and oxygen atoms are known as the main chain or protein backbone.[25]: 19 

    The peptide bond has two resonance forms that contribute some double-bond character and inhibit rotation around its axis, so that the alpha carbons are roughly coplanar. The other two dihedral angles in the peptide bond determine the local shape assumed by the protein backbone.[25]: 31  The end with a free amino group is known as the N-terminus or amino terminus, whereas the end of the protein with a free carboxyl group is known as the C-terminus or carboxy terminus (the sequence of the protein is written from N-terminus to C-terminus, from left to right).

    The words protein, polypeptide, and peptide are a little ambiguous and can overlap in meaning. Protein is generally used to refer to the complete biological molecule in a stable conformation, whereas peptide is generally reserved for a short amino acid oligomers often lacking a stable 3D structure. But the boundary between the two is not well defined and usually lies near 20–30 residues.[26]Polypeptide can refer to any single linear chain of amino acids, usually regardless of length, but often implies an absence of a defined conformation.


    Proteins can interact with many types of molecules, including with other proteins, with lipids, with carboyhydrates, and with DNA.[27][28][25][29]

    Abundance in cells

    It has been estimated that average-sized bacteria contain about 2 million proteins per cell (e.g. E. coli and Staphylococcus aureus). Smaller bacteria, such as Mycoplasma or spirochetes contain fewer molecules, on the order of 50,000 to 1 million. By contrast, eukaryotic cells are larger and thus contain much more protein. For instance, yeast cells have been estimated to contain about 50 million proteins and human cells on the order of 1 to 3 billion.[30] The concentration of individual protein copies ranges from a few molecules per cell up to 20 million.[31] Not all genes coding proteins are expressed in most cells and their number depends on, for example, cell type and external stimuli. For instance, of the 20,000 or so proteins encoded by the human genome, only 6,000 are detected in lymphoblastoid cells.[32]



    A ribosome produces a protein using mRNA as template
    The DNAsequence of a gene encodesthe amino acid sequence of a protein

    Main article: Protein biosynthesis

    Proteins are assembled from amino acids using information encoded in genes. Each protein has its own unique amino acid sequence that is specified by the nucleotide sequence of the gene encoding this protein. The genetic code is a set of three-nucleotide sets called codons and each three-nucleotide combination designates an amino acid, for example AUG (adenine–uracil–guanine) is the code for methionine. Because DNA contains four nucleotides, the total number of possible codons is 64; hence, there is some redundancy in the genetic code, with some amino acids specified by more than one codon.[29]: 1002–42  Genes encoded in DNA are first transcribed into pre-messenger RNA (mRNA) by proteins such as RNA polymerase. Most organisms then process the pre-mRNA (also known as a primary transcript) using various forms of Post-transcriptional modification to form the mature mRNA, which is then used as a template for protein synthesis by the ribosome. In prokaryotes the mRNA may either be used as soon as it is produced, or be bound by a ribosome after having moved away from the nucleoid. In contrast, eukaryotes make mRNA in the cell nucleus and then translocate it across the nuclear membrane into the cytoplasm, where protein synthesis then takes place. The rate of protein synthesis is higher in prokaryotes than eukaryotes and can reach up to 20 amino acids per second.[33]

    The process of synthesizing a protein from an mRNA template is known as translation. The mRNA is loaded onto the ribosome and is read three nucleotides at a time by matching each codon to its base pairinganticodon located on a transfer RNA molecule, which carries the amino acid corresponding to the codon it recognizes. The enzyme aminoacyl tRNA synthetase "charges" the tRNA molecules with the correct amino acids. The growing polypeptide is often termed the nascent chain. Proteins are always biosynthesized from N-terminus to C-terminus.[29]: 1002–42 

    The size of a synthesized protein can be measured by the number of amino acids it contains and by its total molecular mass, which is normally reported in units of daltons (synonymous with atomic mass units), or the derivative unit kilodalton (kDa). The average size of a protein increases from Archaea to Bacteria to Eukaryote (283, 311, 438 residues and 31, 34, 49 kDa respectively) due to a bigger number of protein domains constituting proteins in higher organisms.[34] For instance, yeast proteins are on average 466 amino acids long and 53 kDa in mass.[26] The largest known proteins are the titins, a component of the musclesarcomere, with a molecular mass of almost 3,000 kDa and a total length of almost 27,000 amino acids.[35]

    Chemical synthesis

    Main article: Peptide synthesis

    Short proteins can also be synthesized chemically by a family of methods known as peptide synthesis, which rely on organic synthesis techniques such as chemical ligation to produce peptides in high yield.[36] Chemical synthesis allows for the introduction of non-natural amino acids into polypeptide chains, such as attachment of fluorescent probes to amino acid side chains.[37] These methods are useful in laboratory biochemistry and cell biology, though generally not for commercial applications. Chemical synthesis is inefficient for polypeptides longer than about 300 amino acids, and the synthesized proteins may not readily assume their native tertiary structure. Most chemical synthesis methods proceed from C-terminus to N-terminus, opposite the biological reaction.[38]


    The crystal structure of the chaperonin, a huge protein complex. A single protein subunit is highlighted. Chaperonins assist protein folding.
    Three possible representations of the three-dimensional structure of the protein triose phosphate isomerase. Left: All-atom representation colored by atom type. Middle:Simplified representation illustrating the backbone conformation, colored by secondary structure. Right: Solvent-accessible surface representation colored by residue type (acidic residues red, basic residues blue, polar residues green, nonpolar residues white).

    Main article: Protein structure

    Further information: Protein structure prediction

    Most proteins fold into unique 3D structures. The shape into which a protein naturally folds is known as its native conformation.[25]: 36  Although many proteins can fold unassisted, simply through the chemical properties of their amino acids, others require the aid of molecular chaperones to fold into their native states.[25]: 37  Biochemists often refer to four distinct aspects of a protein's structure:[25]: 30–34 

    • Primary structure: the amino acid sequence. A protein is a polyamide.
    • Secondary structure: regularly repeating local structures stabilized by hydrogen bonds. The most common examples are the α-helix, β-sheet and turns. Because secondary structures are local, many regions of different secondary structure can be present in the same protein molecule.
    • Tertiary structure: the overall shape of a single protein molecule; the spatial relationship of the secondary structures to one another. Tertiary structure is generally stabilized by nonlocal interactions, most commonly the formation of a hydrophobic core, but also through salt bridges, hydrogen bonds, disulfide bonds, and even posttranslational modifications. The term "tertiary structure" is often used as synonymous with the term fold. The tertiary structure is what controls the basic function of the protein.
    • Quaternary structure: the structure formed by several protein molecules (polypeptide chains), usually called protein subunits in this context, which function as a single protein complex.
    • Quinary structure: the signatures of protein surface that organize the crowded cellular interior. Quinary structure is dependent on transient, yet essential, macromolecular interactions that occur inside living cells.

    Proteins are not entirely rigid molecules. In addition to these levels of structure, proteins may shift between several related structures while they perform their functions. In the context of these functional rearrangements, these tertiary or quaternary structures are usually referred to as "conformations", and transitions between them are called conformational changes. Such changes are often induced by the binding of a substrate molecule to an enzyme's active site, or the physical region of the protein that participates in chemical catalysis. In solution proteins also undergo variation in structure through thermal vibration and the collision with other molecules.[29]: 368–75 

    Proteins can be informally divided into three main classes, which correlate with typical tertiary structures: globular proteins, fibrous proteins, and membrane proteins. Almost all globular proteins are soluble and many are enzymes. Fibrous proteins are often structural, such as collagen, the major component of connective tissue, or keratin, the protein component of hair and nails. Membrane proteins often serve as receptors or provide channels for polar or charged molecules to pass through the cell membrane.[29]: 165–85 

    A special case of intramolecular hydrogen bonds within proteins, poorly shielded from water attack and hence promoting their own dehydration, are called dehydrons.[39]

    Protein domains

    Main article: Protein domain

    Many proteins are composed of several protein domains, i.e. segments of a protein that fold into distinct structural units. Domains usually also have specific functions, such as enzymatic activities (e.g. kinase) or they serve as binding modules (e.g. the SH3 domain binds to proline-rich sequences in other proteins).

    Sequence motif

    Short amino acid sequences within proteins often act as recognition sites for other proteins.[40] For instance, SH3 domains typically bind to short PxxP motifs (i.e. 2 prolines [P], separated by two unspecified amino acids [x], although the surrounding amino acids may determine the exact binding specificity). Many such motifs has been collected in the Eukaryotic Linear Motif (ELM) database.

    Cellular functions

    Proteins are the chief actors within the cell, said to be carrying out the duties specified by the information encoded in genes.[26] With the exception of certain types of RNA, most other biological molecules are relatively inert elements upon which proteins act. Proteins make up half the dry weight of an Escherichia coli cell, whereas other macromolecules such as DNA and RNA make up only 3% and 20%, respectively.[41] The set of proteins expressed in a particular cell or cell type is known as its proteome.

    The enzyme hexokinaseis shown as a conventional ball-and-stick molecular model. To scale in the top right-hand corner are two of its substrates, ATPand glucose.

    The chief characteristic of proteins that also allows their diverse set of functions is their ability to bind other molecules specifically and tightly. The region of the protein responsible for binding another molecule is known as the binding site and is often a depression or "pocket" on the molecular surface. This binding ability is mediated by the tertiary structure of the protein, which defines the binding site pocket, and by the chemical properties of the surrounding amino acids' side chains. Protein binding can be extraordinarily tight and specific; for example, the ribonuclease inhibitor protein binds to human angiogenin with a sub-femtomolar dissociation constant (<10−15 M) but does not bind at all to its amphibian homolog onconase (>1 M). Extremely minor chemical changes such as the addition of a single methyl group to a binding partner can sometimes suffice to nearly eliminate binding; for example, the aminoacyl tRNA synthetase specific to the amino acid valine discriminates against the very similar side chain of the amino acid isoleucine.[42]

    Proteins can bind to other proteins as well as to small-molecule substrates. When proteins bind specifically to other copies of the same molecule, they can oligomerize to form fibrils; this process occurs often in structural proteins that consist of globular monomers that self-associate to form rigid fibers. Protein–protein interactions also regulate enzymatic activity, control progression through the cell cycle, and allow the assembly of large protein complexes that carry out many closely related reactions with a common biological function. Proteins can also bind to, or even be integrated into, cell membranes. The ability of binding partners to induce conformational changes in proteins allows the construction of enormously complex signaling networks.[29]: 830–49  As interactions between proteins are reversible, and depend heavily on the availability of different groups of partner proteins to form aggregates that are capable to carry out discrete sets of function, study of the interactions between specific proteins is a key to understand important aspects of cellular function, and ultimately the properties that distinguish particular cell types.[43][44]


    Main article: Enzyme

    The best-known role of proteins in the cell is as enzymes, which catalyse chemical reactions. Enzymes are usually highly specific and accelerate only one or a few chemical reactions. Enzymes carry out most of the reactions involved in metabolism, as well as manipulating DNA in processes such as DNA replication, DNA repair, and transcription. Some enzymes act on other proteins to add or remove chemical groups in a process known as posttranslational modification. About 4,000 reactions are known to be catalysed by enzymes.[45] The rate acceleration conferred by enzymatic catalysis is often enormous—as much as 1017-fold increase in rate over the uncatalysed reaction in the case of orotate decarboxylase (78 million years without the enzyme, 18 milliseconds with the enzyme).[46]

    The molecules bound and acted upon by enzymes are called substrates. Although enzymes can consist of hundreds of amino acids, it is usually only a small fraction of the residues that come in contact with the substrate, and an even smaller fraction—three to four residues on average—that are directly involved in catalysis.[47] The region of the enzyme that binds the substrate and contains the catalytic residues is known as the active site.

    Dirigent proteins are members of a class of proteins that dictate the stereochemistry of a compound synthesized by other enzymes.[48]

    Cell signaling and ligand binding

    See also: Glycan-protein interactions

    Many proteins are involved in the process of cell signaling and signal transduction. Some proteins, such as insulin, are extracellular proteins that transmit a signal from the cell in which they were synthesized to other cells in distant tissues. Others are membrane proteins that act as receptors whose main function is to bind a signaling molecule and induce a biochemical response in the cell. Many receptors have a binding site exposed on the cell surface and an effector domain within the cell, which may have enzymatic activity or may undergo a conformational change detected by other proteins within the cell.[28]: 251–81 

    Antibodies are protein components of an adaptive immune system whose main function is to bind antigens, or foreign substances in the body, and target them for destruction. Antibodies can be secreted into the extracellular environment or anchored in the membranes of specialized B cells known as plasma cells. Whereas enzymes are limited in their binding affinity for their substrates by the necessity of conducting their reaction, antibodies have no such constraints. An antibody's binding affinity to its target is extraordinarily high.[29]: 275–50 

    Many ligand transport proteins bind particular small biomolecules and transport them to other locations in the body of a multicellular organism. These proteins must have a high binding affinity when their ligand is present in high concentrations, but must also release the ligand when it is present at low concentrations in the target tissues. The canonical example of a ligand-binding protein is haemoglobin, which transports oxygen from the lungs to other organs and tissues in all vertebrates and has close homologs in every biological kingdom.[29]: 222–29 Lectins are sugar-binding proteins which are highly specific for their sugar moieties. Lectins typically play a role in biological recognition phenomena involving cells and proteins.[49]Receptors and hormones are highly specific binding proteins.

    Transmembrane proteins can also serve as ligand transport proteins that alter the permeability of the cell membrane to small molecules and ions. The membrane alone has a hydrophobic core through which polar or charged molecules cannot diffuse. Membrane proteins contain internal channels that allow such molecules to enter and exit the cell. Many ion channel proteins are specialized to select for only a particular ion; for example, potassium and sodium channels often discriminate for only one of the two ions.[28]: 232–34 

    Structural proteins

    Structural proteins confer stiffness and rigidity to otherwise-fluid biological components. Most structural proteins are fibrous proteins; for example, collagen and elastin are critical components of connective tissue such as cartilage, and keratin is found in hard or filamentous structures such as hair, nails, feathers, hooves, and some animal shells.[29]: 178–81  Some globular proteins can also play structural functions, for example, actin and tubulin are globular and soluble as monomers, but polymerize to form long, stiff fibers that make up the cytoskeleton, which allows the cell to maintain its shape and size.

    Other proteins that serve structural functions are motor proteins such as myosin, kinesin, and dynein, which are capable of generating mechanical forces. These proteins are crucial for cellular motility of single celled organisms and the sperm of many multicellular organisms which reproduce sexually. They also generate the forces exerted by contracting muscles[29]: 258–64, 272  and play essential roles in intracellular transport.

    Protein evolution

    Main article: Molecular evolution

    A key question in molecular biology is how proteins evolve, i.e. how can mutations (or rather changes in amino acid sequence) lead to new structures and functions? Most amino acids in a protein can be changed without disrupting activity or function, as can be seen from numerous homologous proteins across species (as collected in specialized databases for protein families, e.g. PFAM).[50] In order to prevent dramatic consequences of mutations, a gene may be duplicated before it can mutate freely. However, this can also lead to complete loss of gene function and thus pseudo-genes.[51] More commonly, single amino acid changes have limited consequences although some can change protein function substantially, especially in enzymes. For instance, many enzymes can change their substrate specificity by one or a few mutations.[52] Changes in substrate specificity are facilitated by substrate promiscuity, i.e. the ability of many enzymes to bind and process multiple substrates. When mutations occur, the specificity of an enzyme can increase (or decrease) and thus its enzymatic activity.[52] Thus, bacteria (or other organisms) can adapt to different food sources, including unnatural substrates such as plastic.[53]

    Methods of study

    Main article: Protein methods

    The activities and structures of proteins may be examined in vitro,in vivo, and in silico. In vitro studies of purified proteins in controlled environments are useful for learning how a protein carries out its function: for example, enzyme kinetics studies explore the chemical mechanism of an enzyme's catalytic activity and its relative affinity for various possible substrate molecules. By contrast, in vivo experiments can provide information about the physiological role of a protein in the context of a cell or even a whole organism. In silico studies use computational methods to study proteins.

    Protein purification

    Main article: Protein purification

    To perform in vitro analysis, a protein must be purified away from other cellular components. This process usually begins with cell lysis, in which a cell's membrane is disrupted and its internal contents released into a solution known as a crude lysate. The resulting mixture can be purified using ultracentrifugation, which fractionates the various cellular components into fractions containing soluble proteins; membrane lipids and proteins; cellular organelles, and nucleic acids. Precipitation by a method known as salting out can concentrate the proteins from this lysate. Various types of chromatography are then used to isolate the protein or proteins of interest based on properties such as molecular weight, net charge and binding affinity.[25]: 21–24  The level of purification can be monitored using various types of gel electrophoresis if the desired protein's molecular weight and isoelectric point are known, by spectroscopy if the protein has distinguishable spectroscopic features, or by enzyme assays if the protein has enzymatic activity. Additionally, proteins can be isolated according to their charge using electrofocusing.[54]

    For natural proteins, a series of purification steps may be necessary to obtain protein sufficiently pure for laboratory applications. To simplify this process, genetic engineering is often used to add chemical features to proteins that make them easier to purify without affecting their structure or activity. Here, a "tag" consisting of a specific amino acid sequence, often a series of histidine residues (a "His-tag"), is attached to one terminus of the protein. As a result, when the lysate is passed over a chromatography column containing nickel, the histidine residues ligate the nickel and attach to the column while the untagged components of the lysate pass unimpeded. A number of different tags have been developed to help researchers purify specific proteins from complex mixtures.[55]

    Cellular localization

    The study of proteins in vivo is often concerned with the synthesis and localization of the protein within the cell. Although many intracellular proteins are synthesized in the cytoplasm and membrane-bound or secreted proteins in the endoplasmic reticulum, the specifics of how proteins are targeted to specific organelles or cellular structures is often unclear. A useful technique for assessing cellular localization uses genetic engineering to express in a cell a fusion protein or chimera consisting of the natural protein of interest linked to a "reporter" such as green fluorescent protein (GFP).[56] The fused protein's position within the cell can be cleanly and efficiently visualized using microscopy,[57] as shown in the figure opposite.

    Other methods for elucidating the cellular location of proteins requires the use of known compartmental markers for regions such as the ER, the Golgi, lysosomes or vacuoles, mitochondria, chloroplasts, plasma membrane, etc. With the use of fluorescently tagged versions of these markers or of antibodies to known markers, it becomes much simpler to identify the localization of a protein of interest. For example, indirect immunofluorescence will allow for fluorescence colocalization and demonstration of location. Fluorescent dyes are used to label cellular compartments for a similar purpose.[58]

    Other possibilities exist, as well. For example, immunohistochemistry usually utilizes an antibody to one or more proteins of interest that are conjugated to enzymes yielding either luminescent or chromogenic signals that can be compared between samples, allowing for localization information. Another applicable technique is cofractionation in sucrose (or other material) gradients using isopycnic centrifugation.[59] While this technique does not prove colocalization of a compartment of known density and the protein of interest, it does increase the likelihood, and is more amenable to large-scale studies.

    Finally, the gold-standard method of cellular localization is immunoelectron microscopy. This technique also uses an antibody to the protein of interest, along with classical electron microscopy techniques. The sample is prepared for normal electron microscopic examination, and then treated with an antibody to the protein of interest that is conjugated to an extremely electro-dense material, usually gold. This allows for the localization of both ultrastructural details as well as the protein of interest.[60]

    Through another genetic engineering application known as site-directed mutagenesis, researchers can alter the protein sequence and hence its structure, cellular localization, and susceptibility to regulation. This technique even allows the incorporation of unnatural amino acids into proteins, using modified tRNAs,[61] and may allow the rational design of new proteins with novel properties.[62]


    Main article: Proteomics

    The total complement of proteins present at a time in a cell or cell type is known as its proteome, and the study of such large-scale data sets defines the field of proteomics, named by analogy to the related field of genomics. Key experimental techniques in proteomics include 2D electrophoresis,[63] which allows the separation of many proteins, mass spectrometry,[64] which allows rapid high-throughput identification of proteins and sequencing of peptides (most often after in-gel digestion), protein microarrays, which allow the detection of the relative levels of the various proteins present in a cell, and two-hybrid screening, which allows the systematic exploration of protein–protein interactions.[65] The total complement of biologically possible such interactions is known as the interactome.[66] A systematic attempt to determine the structures of proteins representing every possible fold is known as structural genomics.[67]

    Structure determination

    Discovering the tertiary structure of a protein, or the quaternary structure of its complexes, can provide important clues about how the protein performs its function and how it can be affected, i.e. in drug design. As proteins are too small to be seen under a light microscope, other methods have to be employed to determine their structure. Common experimental methods include X-ray crystallography and NMR spectroscopy, both of which can produce structural information at atomic resolution. However, NMR experiments are able to provide information from which a subset of distances between pairs of atoms can be estimated, and the final possible conformations for a protein are determined by solving a distance geometry problem. Dual polarisation interferometry is a quantitative analytical method for measuring the overall protein conformation and conformational changes due to interactions or other stimulus. Circular dichroism is another laboratory technique for determining internal β-sheet / α-helical composition of proteins. Cryoelectron microscopy is used to produce lower-resolution structural information about very large protein complexes, including assembled viruses;[28]: 340–41  a variant known as electron crystallography can also produce high-resolution information in some cases, especially for two-dimensional crystals of membrane proteins.[68] Solved structures are usually deposited in the Protein Data Bank (PDB), a freely available resource from which structural data about thousands of proteins can be obtained in the form of Cartesian coordinates for each atom in the protein.[69]

    Many more gene sequences are known than protein structures. Further, the set of solved structures is biased toward proteins that can be easily subjected to the conditions required in X-ray crystallography, one of the major structure determination methods. In particular, globular proteins are comparatively easy to crystallize in preparation for X-ray crystallography. Membrane proteins and large protein complexes, by contrast, are difficult to crystallize and are underrepresented in the PDB.[70]Structural genomics initiatives have attempted to remedy these deficiencies by systematically solving representative structures of major fold classes. Protein structure prediction methods attempt to provide a means of generating a plausible structure for proteins whose structures have not been experimentally determined.[71]

    Structure prediction

    Constituent amino-acids can be analyzed to predict secondary, tertiary and quaternary protein structure, in this case hemoglobin containing hemeunits

    Main articles: Protein structure prediction and List of protein structure prediction software

    Complementary to the field of structural genomics, protein structure prediction develops efficient mathematical models of proteins to computationally predict the molecular formations in theory, instead of detecting structures with laboratory observation.[72] The most successful type of structure prediction, known as homology modeling, relies on the existence of a "template" structure with sequence similarity to the protein being modeled; structural genomics' goal is to provide sufficient representation in solved structures to model most of those that remain.[73] Although producing accurate models remains a challenge when only distantly related template structures are available, it has been suggested that sequence alignment is the bottleneck in this process, as quite accurate models can be produced if a "perfect" sequence alignment is known.[74] Many structure prediction methods have served to inform the emerging field of protein engineering, in which novel protein folds have already been designed.[75] Also proteins (in eukaryotes ~33%) contain large unstructured but biologically functional segments and can be classified as intrinsically disordered proteins.[76] Predicting and analysing protein disorder is, therefore, an important part of protein structure characterisation.[77]


    Main article: Bioinformatics

    A vast array of computational methods have been developed to analyze the structure, function and evolution of proteins. The development of such tools has been driven by the large amount of genomic and proteomic data available for a variety of organisms, including the human genome. It is simply impossible to study all proteins experimentally, hence only a few are subjected to laboratory experiments while computational tools are used to extrapolate to similar proteins. Such homologous proteins can be efficiently identified in distantly related organisms by sequence alignment. Genome and gene sequences can be searched by a variety of tools for certain properties. Sequence profiling tools can find restriction enzyme sites, open reading frames in nucleotide sequences, and predict secondary structures. Phylogenetic trees can be constructed and evolutionary hypotheses developed using special software like ClustalW regarding the ancestry of modern organisms and the genes they express. The field of bioinformatics is now indispensable for the analysis of genes and proteins.

    In silico simulation of dynamical processes

    A more complex computational problem is the prediction of intermolecular interactions, such as in molecular docking,[78]protein folding, protein–protein interaction and chemical reactivity. Mathematical models to simulate these dynamical processes involve molecular mechanics, in particular, molecular dynamics. In this regard, in silico simulations discovered the folding of small α-helical protein domains such as the villin headpiece,[79] the HIV accessory protein[80] and hybrid methods combining standard molecular dynamics with quantum mechanical mathematics have explored the electronic states of rhodopsins.[81]

    Beyond classical molecular dynamics, quantum dynamics methods allow the simulation of proteins in atomistic detail with an accurate description of quantum mechanical effects. Examples include the multi-layer multi-configuration time-dependent Hartree (MCTDH) method and the hierarchical equations of motion (HEOM) approach, which have been applied to plant cryptochromes[82] and bacteria light-harvesting complexes,[83] respectively. Both quantum and classical mechanical simulations of biological-scale systems are extremely computationally demanding, so distributed computing initiatives (for example, the [email protected] project[84]) facilitate the molecular modeling by exploiting advances in GPU parallel processing and Monte Carlo techniques.

    Chemical analysis

    The total nitrogen content of organic matter is mainly formed by the amino groups in proteins. The Total Kjeldahl Nitrogen (TKN) is a measure of nitrogen widely used in the analysis of (waste) water, soil, food, feed and organic matter in general. As the name suggests, the Kjeldahl method is applied. More sensitive methods are available.[85][86]


    Further information: Protein (nutrient) and Protein quality

    Most microorganisms and plants can biosynthesize all 20 standard amino acids, while animals (including humans) must obtain some of the amino acids from the diet.[41] The amino acids that an organism cannot synthesize on its own are referred to as essential amino acids. Key enzymes that synthesize certain amino acids are not present in animals—such as aspartokinase, which catalyses the first step in the synthesis of lysine, methionine, and threonine from aspartate. If amino acids are present in the environment, microorganisms can conserve energy by taking up the amino acids from their surroundings and downregulating their biosynthetic pathways.

    In animals, amino acids are obtained through the consumption of foods containing protein. Ingested proteins are then broken down into amino acids through digestion, which typically involves denaturation of the protein through exposure to acid and hydrolysis by enzymes called proteases. Some ingested amino acids are used for protein biosynthesis, while others are converted to glucose through gluconeogenesis, or fed into the citric acid cycle. This use of protein as a fuel is particularly important under starvation conditions as it allows the body's own proteins to be used to support life, particularly those found in muscle.[87]

    In animals such as dogs and cats, protein maintains the health and quality of the skin by promoting hair follicle growth and keratinization, and thus reducing the likelihood of skin problems producing malodours.[88] Poor-quality proteins also have a role regarding gastrointestinal health, increasing the potential for flatulence and odorous compounds in dogs because when proteins reach the colon in an undigested state, they are fermented producing hydrogen sulfide gas, indole, and skatole.[89] Dogs and cats digest animal proteins better than those from plants, but products of low-quality animal origin are poorly digested, including skin, feathers, and connective tissue.[89]

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