Chemical Modifications of Biotech Drugs PDF

CHEMICAL MODIFICATIONS OF BIOTECH DRUGS Alternative approaches to overcome the degradation of peptides, proteins and oligonucleotides could be of replacement of those aminoacidic residues that are sites of proteolytic degradation by genetic engineering, of synthesis of truncated and more stable prot...

CHEMICAL MODIFICATIONS OF BIOTECH DRUGS Alternative approaches to overcome the degradation of peptides, proteins and oligonucleotides could be of replacement of those aminoacidic residues that are sites of proteolytic degradation by genetic engineering, of synthesis of truncated and more stable protein sequence, of entrapment into particles such as liposomes or insoluble polymeric microspheres or of covalent linking of the surface polymers. This last technique involves the use of either natural or synthetic polymers, but the main used is the PEG. Polyethylene glycol is a molecule that presents only one reactive terminal group per polymer chain and does not cross-link or react with any other molecule. The purpose of this molecule, like for every other polymer added to biotech drugs, is to prevent the proteins being recognized by the proteolytic enzymes and by antibodies and to lower the positive charge of the cationic complexes for nucleic-based drugs. In fact with the PEGylation we prolong the half-life of our drug (especially for proteins). PEG could also be used to link together two variable portions of an antibody molecule, an example could be the anti-CD4 PEG-Fab molecule, with PEG that conjugates the two Fab portions recognizing the CD4 by the hinging disulphides (peptides, proteins, enzymes and antibody fragments can be site-specifically PEGylated using a native and accessible disulfide without destroying the molecules' tertiary structure or abolishing its biological activity). So, the advantages of PEG as drug carrier are: increased retention time in blood as a consequence of a reduced glomerular filtration (because PEG is a big molecule; this is particular evident for low molecular weight compounds such as peptides and oligonucleotides); surface masking of the drug to avoid proteolysis and binding to antibodies; the possibility of linking the drug to one end of the PEG chain and a targeting agent at the opposite end (so to direct to target sites in the body); PEG creates a hydrophilic cloud around the molecule. BIOTECH DRUGS: GENE MANIPULATION AND RECOMBINANT DNA TECHNOLOGY The protein that we have to use as a drug has to be produced by a gene, so we first have to clone the gene that encodes for our therapeutic protein inside an expression vector. This will then be inserted in our host cell, that is going to produce the protein that we want to use as a drug. So let’s start with the cloning and preparation of the expression vector: first you select the gene of the protein that you want to use as a drug, you amplify the gene with a PCR and then you insert it in a vector through the use of a ligase. The vector that you choose for the expression has to be compatible with the host cell in which you want to produce your protein, meaning that you have to insert, other than the gene, also controlling sequences that are typical of the expression of proteins in that specie. So in the vector we have to insert also a promoter, with a transcription starting point, and a transcriptional termination site. For example, signals that ensure gene expression in procaryotes are distinct, and these signals must be contained in the vector if we chose to produce our therapeutic protein in bacteria, because when a eukaryotic gene is transferred to a bacterial cell, this gene has a low chance to be expressed. So to be sure that a correct transcription and translation of foreign genes can take place in the host cell and that the protein, after its expression, will not get digested because it is recognized as exogenous, we need to pay attention to that. To obtain the maximum yield, it is necessary to optimize the stability of the products, both at the transcriptional and translational levels. The optimization of transcription is mainly operated through the regulation sequences, especially from the promoter. The expression level of a gene depends largely on the strength of the promoter, which determines the frequency with which the RNA polymerase starts the transcription. Since the vectors that we use are mainly prokaryotic vectors (in fact they are plasmids) and since the expression usually happens in bacterial cells, many strong promoters of E. coli have been isolated and inserted in the available expression vectors, however some synthetic promoter have also been produced on the basis of optimal consensus sequences for a specific expression. A typical procaryotic promoter contain about 60 bp with 2 consensus sequences located are -35 (TTCAGA) and -10 (TATAAT); some regions called “up” have been recently characterized to enhance the efficiency of transcription. The most common promoters are the Lac, lacUV10, trc, tac, T3, T7, Lambda Pr, Lambda Pl. Also the end of the transcription of the protein follows a specific signal which are termination signals. These signals differ between eukaryotes and prokaryotes, the ones that bacteria use are generally of two types: rho-independent (also known as intrinsic termination, it relies on the formation of a GC-rich hairpin in the RNA transcript followed by a weakly bound poly-uracil tract) and rho-dependent (they are not usually employed in the plasmid-based expression systems, they depend on three factors, which are rho, tau and NusA). So in general the termination of the transcription could occur because of the formation of tertiary structured in the mRNA or because of the recruitment of some enzymes, but the one that happens with the formation of a tertiary folding is the preferred one for this applications. The optimization of translation is achieved thanks to the sequence of SHINE-DALGARNO (5’-AGGAGGU-3’), a binding region for the ribosome in the 5’ untranslated region of the mRNA. Next to the Shine-Dalgarno sequence a start codon must be found, most of the time AUG (sometimes also GUG); the three bases that precede AUG are important for the stability of the protein, which can vary up to 20% when these bases are changed. The optimal number of bases between Shine-Dalgarno and AUG is 8 bases. What is mostly important is the optimization of the stability of the protein inside our host cell: because the yield of an expression product depends mainly on the stability of the protein, and the stability of the protein depends on the presence of stabilizing amino acids at the N-terminal and on the de-stabilizing amino acids at the C-terminal of the protein. For example, the beta-galactosidase stability depends on the amino acids at the N-terminal of the protein: if we have either methionine, serine, alanine, threonine, valine or glycine, the half-life lasts for more than 20 hours, but if we arginine, the half-life could last only 1 minute! This is because PEST sequences (rich in proline (P), glutamic acid (E), serin (S) and threonine (T) and flanked by positively charged AA), are target of degradation and make bacterial proteins less stable. This suggests that the modification of the codifying sequence through genetic engineering could lead to a modified aminoacidic sequence that may have a better stability than the original sequence of the protein (for example, removing the PEST sequences by site-specific mutagenesis, can improve the stability of recombinant proteins). However, aminoacidic sequences cannot be altered in therapeutic proteins, otherwise the tertiary structure of the protein, and thus its function, could be altered! So if we want to improve the stability of protein, for example removing the PEST sequences, we have to make sure that the protein thus produced will still be effective in its therapeutic activity. Other strategies that could be used to optimize the translation grade and the stability of the protein are to use alternative codons (synonymous codons, so codons that can generate the same amino acid, but that increase the transcript’s level of expression without modifying the expression product) or to use protease-free species (we reduce the proteolytic degradation inside the host cell so that our exogenous protein won’t be a target of these enzymes; the presence of more than 20 proteases in a bacterial cell makes this task difficult, however there are some mutant E. coli species with defective proteases, like ompT; this strategy is used to increase the stability of the protein when we cannot change the sequence of the aminoacidic chain of the protein otherwise we will alter its function). However, expression vectors also need to be controlled: a continuous production of a protein could kill bacterial cells, causing inhibition of cell functions, loss of energy, loss of plasmid. To avoid this problem we can use specific vectors in which we induce the transcription of the protein just in a specific moment. These vectors are called inducible vectors and they work associated to substances which are able to induce their expression only in the case in which they are present. In this way, the bacterial system doesn’t overload and we are able to achieve an optimal production in a limited period of time, called induction phase. If we don’t use these strategy and we, for example, leave the bacteria grow in two days, it is possible that our protein will be produced only for the first two hours and then it won’t be produced anymore, either because the protein that we want the bacterium to express can be toxic for the host cell, or because, under some conditions, the recombinant protein can account for up to 30-40% of the total proteins, replacing those that are important for the metabolism of the bacterium. Moreover, when the protein is “induced”, proteolytic degradations are limited and the yield of production of the protein is improved. Another strategy of modification is to produce the protein in a specific region of the bacterial cell or directly outside of it. This could be performed to avoid the degradation of the heterologous protein inside the cell and to facilitate the purification process. The localizations could be: intracellular, periplasmatic, cell wall, extracellular. Except for the intracellular one, all the other localizations could induce a secretion of the protein so that we have the final product directly in the medium and it will be easier to recollect it. The eventual disulphide bonds of the protein occur during secretion, the N-terminal region will be identical to the original product and the correct folding is also guaranteed. All of these can be achieved by producing fusion proteins: for example, we can insert in the vector the gene of our protein of interest fused with the gene for another protein (for example a maltose binding protein), the product will be a fusion protein that can be used to achieve a different localization of the protein in the host cell or to achieve a better purification step (for example, in this case we exploit maltose bound to a column and we make our fusion protein bind to it). Then, the protein used for the purification will be removed and we will simply obtain our therapeutic protein. So in general fusion proteins can be used to increase the stability of the protein during the expression in a different localization of the cell, or to improve the purification steps. For this last purpose, the GST is one of the most used gene in laboratory practice (check table). BIOTECH DRUGS: PROCESSING AND PRODUCTION Which host cells can be used for the production of biotech drugs? 1. E. COLI: it is very easy to grow, it is cheap, we know all the media and all the techniques that are needed to grow this bacterium, so we have a lot of proteic drug that have been and are still produced inside these cells (insulin, INF-alpha, INF-gamma, IL-2, G-CSF, hGH and tPA). Moreover, the level of expression in E. coli is below 30% of the total proteins of the cell, we cannot obtain a higher percentage of protein, but it is still very high. E. coli cells grow rapidly on simple and inexpensive media, their molecular biology is well characterized and their appropriate fermentation technology is well established, so they are really easy to control for us. In E. coli we can produce potentially any type of protein that doesn’t need to be glycosylated. This is the only main problem of this type of cell: glycosylation cannot happen in bacterial cells, it’s a step that can only happen in eukaryotic cells. Another drawback is that, being bacterial cells, they present the LPS on their cell  wall: LPS is a really dangerous molecule for humans, it triggers the production and release of IL- 1beta (pyrogen), it causes endotoxemia and could potentially lead to the death of the individual. So when we produce proteins in E. coli, during the purification step, we have to be sure that not even one molecule of LPS will be present in the final result. Lastly, heterologous proteins usually accumulate intracellularly, which is a disadvantage because, since bacteria present a cell wall, they are very difficult to break through. So every time we have to purify proteins from E. coli cells we have to use homogenization, centrifugation or filtration to remove the cell debris and extensive chromatographic purifications to separate the protein of interest from the several thousand other proteins of the bacterium. One characteristic of E. coli, which could be a problem for biotech drug production, is that they produce inclusion, or refractile, bodies: these bodies are insoluble aggregates constituted by partially folded heterologous products. They could be taken advantage of because they facilitate the purification of the proteic product by a single centrifugation because of their high density, however they could present a problem because in these inclusion bodies our protein is present in a tertiary folding that is not the functional one. How can we prevent the formation of inclusion bodies? We could use a plasmid that fuses our protein with thioredoxin, a molecule that prevents the formation of inclusion bodies and that could be easily removed once our protein is extracted from the E. coli simply by digestion. Another strategy is the reduction of the growth temperature (from 37° to 30°), which avoids the formation of aggregates; the third strategy is the co-expression with chaperons, proteins that favour the correct folding of the protein, so they avoid the aggregation of our biotech drug (by attracting water molecules). Molecular chaperones in fact usually interact with unfolded or partially folded protein subunits, like nascent chains emerging from the ribosome, or extended chains being translocated across subcellular membranes. They stabilize non-native conformation and facilitate correct folding of protein subunits. Some chaperones are non-specific, and interact with a wide variety of polypeptide chains, but others are restricted to specific targets. They often couple ATP binding/hydrolysis to the folding process; they are essential for viability and their expression is often increased by cellular stress. When a protein is denatured, it becomes stabilized in a new conformation by its interaction with the water molecules surrounding the protein. When certain classes of surfactants are added, their hydrophobic sections are attracted to hydrophobic regions of the protein. This induces the positioning of highly polar parts of the surfactants into the water surrounding the protein, allowing the surfactants to disrupt the water structure, lower surface tension and facilitate refolding. By favoring the correct folding, the aggregation of mis-folded proteins is prevented. If we want to exploit the formation of inclusion bodies for the purification process, after the collection the proteins must be incubated with detergents (like urea), which are then removed by dialysis or diafiltration; in this way the protein refolds in its native and biologically active conformation. 2. MAMMALIAN CELLS (animal cell cultures): the main cell lines used are the CHO (hamster cell lines) and the BHK (baby hamster kidney cells). They are able to glycosylate the protein (which is their main advantage), but they need complex media to grow (because mammalian cells can be infected by bacteria and viruses), they grow really slowly, they are more susceptible to physical damage (because they don’t have a cell wall), so they are more fragile especially in culturing conditions. So because of all of these characteristics, their cultivation is more expensive and these cells have high costs. 3. YEAST CELLS (Saccharomyces cerevisiae): their molecular biology is well known and they allow an easy genetic manipulation. They are classified as GRAS organisms (“generally regarded as safe”, this means that we can eat them without getting infection, however for laboratory practice sometimes they are not safe so we have to pay attention). They rapidly grow in inexpensive medium; their cell wall protects them from physical damage and they are suitable for industrial-scale fermentation. They can carry out post-translational modifications of proteins, included glycosylation, however they carry it out a little differently than mammalian cells, so compared to the glycosylation that we should have in the native protein. The disadvantage, other than these glycosylation problem, is that the expression levels of heterologous human protein remains less than 5% of total cellular proteins.  Despite such disadvantages, several recombinant biopharmaceuticals are produced in yeast (insulin, growth factors, the anticoagulant hirudin), but they are not glycosylated (glycosylation in yeast is made with mannose and it triggers the rapid clearance of the protein from the blood stream and can be immunogenic in humans). 4. FUNGAL PRODUCTION SYSTEM: eukaryotic cells. Suitable for fermentation technology, fungi are capable of high-level expression of various proteins. Moreover, they are able to secrete the proteins into their extracellular media (so you don’t have to extract it) and they can carry out post- transcriptional modifications (even if, also in this case, the pattern of modification can differ from typical patterns in mammalian cell lines). However, until now, no biotech drug produced by such a means has gained marketing approval. 5. TRANSEGNIC ANIMALS: we can produce proteins inside the milk of a transgenic animal (transgenic animal by definition means an animal that is modified to produce a high quantity of a specific protein). This is a great advantage because we know the composition of the milk and so we can easily isolate the produced protein from it. Transgenic animals are considered live bioreactors for the production of proteins of interest. Mammary-specific expression can be achieved by fusing the gene of interest with the promoter containing regulatory sequence of a gene coding for a milk- specific protein (e.g. whey acidic protein -WAP-, b-casein, b-lactoglobulin). As for now, only goats have been used with this specific purpose. They have been engineered to produce tPA (tissue type plasminogen activator) for the treatment of stroke, however this drug has not been marketed yet. Goats usually produce 700-800 L of milk per year. Many other attempts have been made to produce other proteins, also in other animals, like pigs, sheep and rabbit, but they are not available in the market yet. The only approved drug (even if only in the US) produced with this type of technique is ATryn, a recombinant human anti-thrombin. How do we modify the genome of an animal for this purpose? We first clone the gene for our protein in a vector under the control of a specific promoter called WAP promoter (murine whey acid promoter). In this way the promoter allows the expression of our gene only in the mammary gland (so not every cell in the animal will express the gene, but only the mammary glands). For example, the tPA has been expressed in goats through a vector which contains the WAP promoter and then the LAtPA cDNA (it encodes for a point mutated form of tPA that is glycosylated differently resulting in longer acting (LA) tPA) and a SV40 polyA signal (SV40 is a virus that provides the poly A signal telling cells mRNA is ready for expression). The production of a specific biopharmaceutical protein in an animal is called pharming. Recombinant goats are produced starting from the vector insertion: we take a zygote and we insert our gene of interest in it (if possible, directly in the nucleus, if not, in the cytoplasm, then the single plasmids will reach the nucleus). The purpose is to make the gene recombine and integrate with the genome of the zygote: we don’t know how many of them will actually succeed in this, most copies of the vector could be broken down, but some of them may enter the nucleus. Once the plasmid with our gene is inserted in the DNA, the recombinant zygote is implanted in a female goat. These female goat will give birth to the transgene offspring. Once these offspring is old enough to become pregnant it will produce milk with our protein. Obviously, this whole process takes a lot of time, several years need to pass before the mother gives birth to the offspring and before the offspring get old enough to produce milk. Which are the advantages of these system? First of all in goats we have a high milk production capacity (which is not achievable with other animals like rabbits or pigs because they produce a lower amount of milk), goats have an easy handling and breeding, milking systems are already available, the gestation and maturation times are lower than other milking species (like cows, which could produce much higher quantities of milk but in longer times so it would be a disadvantage), high expression levels of proteins can be achieved (>1g/litre milk; in 1 day a goat would produce a similar quantity of product as would be likely recoverable from 50-100 litres in bioreactors). Moreover, as we already said, milk is biochemically well characterized and this helps rational development of appropriate downstream processing protocols. The disadvantages are that in many cases the expression levels are 1mg/L; we always have to characterize the exact nature of the post-translational modifications by the mammary gland; there is a lag time to obtain the  protein (a viable embryo with the inserted gene must be brought to term and this gestation period ranges from 1 month for rabbits to 9 months for cows, after breeding they must bring their offspring to term before they begin to lactate); after some generations of transgenic offspring the recombinant gene is lost, so the animals will stop producing our protein of interest. So the microinjection technique is time-consuming and does not guarantee success. For all these reasons sometimes instead of using this technique we prefer the cloning technique (nuclear transfer): in this case the donor is an adult cell (any tissue cell from an adult goat), you remove the nucleus and you genetically engineer it to harbour the gene of interest, then you substitute it with the nucleus of an egg cells taken from an unfertilized animal. The “reconstructed” embryo is then grown for 7 days, then it is implanted in a surrogate mother and in this way we give birth to a cloned animal with the exact DNA as the tissue cell donor animal. This is the technique that has also been used for the Dolly sheep cloning. The problem with this technique is that the cloned animal could have a short life and health problems. So the microinjection is still preferred in the majority of the cases. In 2014, the first recombinant drug purified from the milk of rabbits was produced: ruconest (International Non-proprietary Name: conestat alfa) is a recombinant human C1 esterase inhibitor (rhC1iNH) developed and approved for the treatment of acute angioedema attacks in patients with hereditary angioedema. HAE is a rare, serious, autosomal-dominant genetic disorder with an estimated prevalence of one in 50,000. Clinically, patients with HAE experience recurrent acute attacks of soft tissue swelling that can affect multiple anatomic regions, including the gastrointestinal tract, facial tissues, vocal cords and larynx, oropharynx, urogenital region, and/or the arms and legs. The transgenic rabbits have been modified to produce the recombinant protein in their milk, however the quantity of milk that we can produce with these animals is not that much. So why do we use rabbits? This disease is very rare, so the amount of protein that we have to produce is not that much, so rabbits are perfect to provide the correct amount of this protein. We only saw the production of biotech drugs in milk, but are there any other biological fluid that could produce recombinant proteins? Potentially, we could use blood, but this means that to purify it we have to make the animal bleed, and this means that the animal could die, so only a low volume of blood can be harvested from animals at any given time, and this is not enough for a pharmaceutical protein production. Moreover, producing proteins in blood means that we could potentially induce side effects, mainly related to immunogenicity, in the producer animal. Another disadvantage is that serum contains a variety of native proteins, so the purification of the recombinant protein could be really complex; in addition, many proteins are poorly stable in serum. So, this production is not suitable for industry. Other fluids where we could think of producing recombinant proteins in transgenic animals are urine and seminal fluid. But these fluids are really difficult to collect, and they are not appropriate for the production of protein in an industrial scale (urine contains toxic molecules and seminal fluid is not produced in high quantities). 4. HEN AS A BIORECTOR: an egg can be easily engineered with retroviruses containing the gene for our biotech protein, to obtain a recombinant egg. Retrovirus particles bearing a transgene, or Barred Rock cESs transformed with an oviduct-specific expression vector, are injected into the sub- germinal cavity of freshly laid, stage X embryos. In the case of retroviruses, the egg is sealed with a plug of hot glue or with eggshell membrane and cement and incubated to hatch, yielding G0 founders chimeric for the transgene. In the case of cESs, the egg is sealed and incubated for 3 days before transfer to a surrogate eggshell and incubated to hatch, yielding high-grade black feathered chimeric chicks. The egg white can contain 4g of proteins, of which 50% is ovalbumin, so eventually just 25% is represented by the recombinant protein. However, once we produce a transgenic rooster, its offspring can produce annually up to 300 eggs, so eventually 300g of recombinant protein can be produced. The main problem consists in the pattern of glycosylation: glycosylation is very different compared to the native glycosylation of human proteins, in fact a lot of attempts have been made with the production of antibodies, but the egg derived mAbs are really immunogenic because of this different glycosylation (reduced serum half-life, of 100 or 200 hours, because the different glycosylation enhances the ADCC activation). For this reason, no biotech drug has been produced with this method yet. 5. TRANSGENIC PLANTS: the advantages of plants for the production of biotech drugs are that plants can be easily grown, the cost for plant cultivation is low, harvest equipment is inexpensive and well established, they are easy to scale-up, pathogens that infect plants cannot infect human beings and they can be easily engineered. However, they have some disadvantages, because sometimes, once you have inserted your transgene in a plant, this can be silenced by intrinsic mechanisms of the plant (post-translational gene silencing); moreover, the glycosylation in plants is completely different from mammalian glycosylation (immunogenicity), the expression levels are often variable, there is an environmental concern for the possible escape of genetically altered plants (for example with pollens) and plants grow in a seasonal/geographical way (not every plant is an evergreen so the production of the proteins should be stopped when the plant stops growing). Agrobacterium tumefaciens and Agrobacterium rhizogenes are the main plants used as vectors. In the future, oral vaccines could be prepared in edible plants (tomatoes, carrots, lettuce, potatoes, rice, tobacco plants and bananas; in any case we can make the protein get expressed both in the leaves and in the products of the plants). However, the percentage of production of proteins in these plants it’s not so high, even if they can be easily engineered. Only one protein has been successfully produced in carrots as for now (2012, FDA approved “Taligurase alfa”, an isoform of glucocerebrosidase, a lysosomal enzyme, that is defective in Gaucher’s disease; produced in carrot cells by Pfizer and Protalix). However, we have to remember that if we produce biotech drugs in plants, we then need to extract the product from the plant, because if we eat the transgenic plant then the protein will be digested! 6. INSECT CELL-BASED SYSTEM: they are derived from silk warms. They have a lot of advantages because they can be easily genetically modified, they can be cultured more rapidly and with less expensive media compared to mammalian cells, they express high levels of proteins, they are not infected by human pathogens; however, again, the main problem is the glycosylation pattern, which is very different from the one of native cells, so we have a high immunogenicity. Moreover, the recombinant protein is usually expressed intracellularly, while the extracellular accumulation of the desired protein is usually low. Some proteins have been obtained in this expression system, like hepatitis B surface antigen, INF-gamma and tPA, but they have all been produced for laboratory practice, so no human protein has been thus far approved and produced in insect cells with therapeutic use (only veterinary vaccines have been produced and approved, against classical swine fever for pigs). The main system used is the baculovirus gene expression in insect cells and silkworm larvae, which can exploit either cells in culture or the whole insect. In the first case the target gene is incorporated into a AcMNPV bacmid (vector). This extracted recombinant AcMNPV bacmid from E. coli is transfected into insect cells, and the resulting recombinant AcMNPV is purified, amplified, and increased in titer and then used for insect cell infection (this technique only requires some weeks). In the second case we have the conventional BmNPV expression system, where the target gene and the wild-type BmNPV gene are co-transfected into B. mori cells, and a recombinant BmNPV is obtained. This recombinant BmNPV bacmid from E. coli is injected directly into silkworm larvae and after 4–6 days post-injection, the recombinant protein is harvested (this technique requires some months, so more time than the other). Comparing all the systems that we just explained, we can see that bacterial cells are the most efficient ones. However, as we talked about, the main problem is glycosylation. In the image we can see, in fact, that the glycosylations that are less immunogenic are the ones performed by mammalian and yeast cells. Biotech drugs production is divided into upstream and downstream processing: the first phase involves the amplification and production of the protein, while the second involves the purification steps and the production of the final pharmaceutical product. 1. UPSTREAM PROCESSING: the first step is to build a cell bank. When a pharmaceutical company comes up with the production of a new pharmaceutical biotechnological product, in the first years the patent policy ensures the owning rights, so only the company will be able to produce and sell it. However, for the production in the company but also for the production by other laboratories after the patent expires, we need to have a specific protocol for the production of the biotech drug that must be always the same, otherwise you obtain a different product. So to be sure that for all the following years the pharmaceutical company will produce the same drug with the same procedure, the company must build a cell bank. So we start with the first culture of the newly constructed production cell line. This culture must be amplified and aliquoted into different vials, which are called master cell bank. Each master cell bank is frozen in liquid nitrogen (-60°/-80°) to preserve it. Once a new production of that drug has to start, one master cell bank will be thawed and it will be amplified and aliquoted in hundreds of vials to produce a working cell bank. These vials of the working cell bank are going to be frozen in turn until they have to be used for the drug production and so they will be thawed (because the production usually happens every n days, like every two weeks for example a vial will be thawed). Each one of these vials of the working cell bank constitutes a batch, so it will have a batch number. This batch number is used to differentiate drugs produced in different producing cycles: we produce biotech drugs at different times starting from different vials, so if something goes wrong with the production from that vial we need to be able, thanks to the batch number, to identify the vial from which the drug was produced. Usually, with biotech drugs, we perform  everything under controlled circumstances, in order to avoid problems with different batches, however there could still be some differences between drug aliquots produced in different producing cycles, for example in quantity. The vials of the cell bank all include frozen cell lines that are able, once thawed, to start a new production of a functional protein. This system is called two- tiered cell banking system. The Master Cell Bank is created because a daily interaction with it would pose an unacceptable risk for any manufacturer (because the majority of the cell line stock will be lost if something goes wrong). For this reason, portions of the MCB are taken-off and stored separately for “day-to-day” use as the feed stock for each production run, soon after creation. These are the working cell banks, and they may be accessed every day, while the remaining stock of the MCB and of the working cell bank are very securely stored away in separate storage tanks. While the WCBs are accessed on a regular basis, the MCB is essentially left untouched as a reserve in the event the WCBs are compromised. Every vial is stored stably in liquid nitrogen at -150 degrees Celsius protecting them over the drug’s entire life cycle. How do we prepare a batch for the production? First, we have the thawing of a vial from the working cell bank, then we seed it in a starter culture, then in a production-scale starter culture, then in a production-scale bioreactor. A bioreactor is a tank that is big enough to contain several thousand or tens of thousands litres of media for the growing of a production-scale cell culture (it is also used for microbial fermentation, in fact the bioreactor can also be called fermenter). So eventually, from a small vial, we obtain litres and litres of production cell cultures. The right temperature for the growth of host cells has to be established prior to this process, and bioreactors allow the control of this parameter. These in the picture are all the steps of the upstream process. How do mammalian cell lines grow in cultures? They require very rich and expensive media, they also require antibiotics and they have to be controlled because they are more fragile than microbial cells due to absence of an outer cell wall. These cells usually secrete the recombinant protein in the medium, so it is easier to recollect it. 2. DOWNSTREAM PROCESS: it mainly consists of the purification process, so it follows the production process. First of all you have to collect the protein, and this step depends on which type of cell line you used for the production process: if you used bacteria you have to break down the cell wall, while if you used mammalian cells you can skip this step and, most of the times, the product recovery is simply performed by harvesting the extracellular fluid (because the biopharmaceuticals are secreted into the media as extracellular proteins). The cell disruption can be performed physically (sonication, homogenization, alkaline conditions exposure, …) or chemically (detergents, solvents, antibiotics and chaotropic agents). However, if you use chemical reagents, then you will find them in the final product so you will have to remove them during purification; moreover, detergents could cause protein denaturation and precipitation. To avoid it, the best choice is to use homogenization (agitation with abrasives), even if also in this case we have to make sure that we don’t use a temperature that is too high, because in that case the proteins can denature. Alternatively, glass beads are used in cellular agitation, but then beads need to be removed, so homogenization is still the best choice. In fact, an efficient cooling system minimizes protein denaturation and inactivation. Upon completion of the homogenization step, cellular debris and the remaining intact cells can be removed by centrifugation or by microfiltration (0,1-10 mm diameter of pores). Then also nucleic acids must be removed (nucleic acids significantly increase the viscosity of the cellular homogenate): nucleic acid content in the final preparation can be, at most, a few pg/therapeutic dose (they are allowed because it is impossible to remove completely nucleic acids from the homogenates). They may be removed by treatment with nucleases, an efficient, inexpensive and innocuous treatment. After the removal of nucleic acids you have to ultrafiltrate the homogenate. For ultrafiltration we use membranes with pore diameters of 1-20 nm so that we can separate molecules based on size and shape. Ultrafiltration doesn’t alter the bioreactivity of the protein, it has high recovery rates (99%) and a short time for recovery and little ancillary equipment is required. In this way, we remove any possible debris that is still contained in the homogenate, because it is going to be retained by the membrane. The proteins of the homogenate will pass the membrane driven by high pressures, they will be eluted and recollected. Instead of using the simple ultrafiltration we could use diafiltration, which is based on the same system but the volume that is inserted in the column of filtration is the same that we collect at the end, because we add the homogenate continuously (so the level of the reservoir is maintained at a constant volume). It is used to reduce or eliminate low molecular mass molecules from a solution (salts, ethanol, buffer components, amino acids, peptides, added protein stabilizers, etc.) and it is generally preceded by an ultrafiltration step. So now we obtained a solution containing only a mixture of the proteins of the cell line, which contains also our protein of interest. Now all contaminants of potential medical significance must be removed, and we do this by chromatography. We have several types of chromatography but there are some specifical types that we use for biotech drugs. Chromatography is based on the characteristics of the specific protein: each protein has a specific size, shape, overall charge, surface hydrophobic groups, … that we define, as a whole, as the chromatographic fingerprint of the protein. In fact, we can differentiate a protein from the rest of the proteic solution on the base of these characteristics. More than one chromatographic type has to be used to obtain a solution containing only our protein of interest, and so a combination of 2 to 4 different chromatographies have to be used in the downstream processing in order to have a highly purified product (at least a level of 99.9% of purity has to be achieved to be sure that, by administering the final product, we don’t have any unwanted reaction; a 100% of purity is not obtainable, so we can just get close to this value). So which types of chromatography can we use in the downstream process? ▪ Size-exclusion chromatography: it is the simplest type because it separates the proteins based on their size and shape. Larger molecules cannot enter the matrix and so they will be quickly eluted, while smaller proteins can enter the gel beads (the smaller the protein, the longer it will be retained in the matrix and so the later it will be eluted). For this reason, if we know that we have to isolate a protein with a high molecular mass we decide to use it at the beginning of the purification process. Instead, when the protein is small, size-exclusion chromatography is employed towards the end of a purification sequence when the protein relatively pure is concentrated in a small volume. Gel matrices that we can use with this technique are for example dextran, agarose, acrylamide or vinyl polymer. Although this chromatography is effective, it results in a significant dilution of the protein solution. ▪ Ion-exchange chromatography: it is the most used chromatographic technique. Some amino acids are negatively charged (aspartate and glutamate), some others are positively charged (arginine, histidine and lysine), so depending on the composition of the protein of interest the protein will have a total positive or negative charge. The charge of proteins depends on the pH of the medium in which they are in solution, and the pH value in which the protein doesn’t have any net charge (positive charges=negative charges) is called isoelectric point (if the pH is below this value, the charge is positive, if the pH is above this value, the charge is negative). The majority of proteins are negatively charged, which means that they contain a higher number of  negatively charged amino acids. So if we use a matrix containing positively charged beads (anionic exchanger), we can separate the positively charged proteins, which will be eluted, from the negatively charged proteins, which will be retained by the matrix. Obviously, the electrostatic attraction that makes the negatively proteins retained in the column, is reversible, so if we change the pH of the buffer of elution (salt- containing buffer) we can change the electrostatic interactions between the proteins and the matrix, making the elution more selective. ▪ Hydrophobic interaction chromatography: some amino acids are hydrophobic, which means that they tend to aggregate together and be surrounded by water molecules. So if we use a matrix on the chromatographic column that is made with hydrophobic groups (phenyl Sepharose gel or octyl Sepharose gel for example), the hydrophobic group of hydrophobic proteins will tend to aggregate with it, while proteins with a less hydrophobic nature will be eluted first (the more hydrophobic the protein, the tighter the binding). Elution is obtained with a buffer of decreased ionic strength. ▪ Affinity chromatography: we use beads made with agarose (or resin), a spacer and a ligand, so a specific protein that is able to bind another specific protein. If for example we need to purify a cytokine (IL-2 for example), we can use, as a ligand in the matrix, a part of the cytokine receptor. Elution can be obtained by changing the pH of the buffer, changing the ionic strength of the buffer, changing the detergent or by using a competing ligand. Affinity chromatography should be used as early as possible in the purification procedure. The main advantage of this type of chromatography is that it is highly specific and selective, and thus the increase in purity is of over 1000-fold. However, many ligands that we usually use in this technique are highly expensive and exhibit a poor stability, and the ligand coupling techniques are chemically complex, hazardous and costly. Moreover, the ligand of the matrix can leach from the column and this can be a problem for the purification. Another type of affinity chromatography is the Abs affinity chromatography: instead of having a protein (like a receptor) has a ligand we use an antibody, which is even more specific for our protein of interest. However the drawbacks are the same as before. Elution is achieved by: altering the pH of the buffer, using chemical disrupting agents such as urea or guanidine, through irrigation with a glycine-HCl buffer (pH 2,2-2,8). This Abs affinity chromatography is used for purification of recombinant blood factor VIII. ▪ Dye affinity chromatography: dyes such as Cibacron blue and Procyon red have been used as adsorbent because of their binding to certain proteins (natural tendency). We know which type of proteins are able to bind to these dyes so we can exploit them for purification. The advantages are that this method is available and inexpensive, the chemical coupling to the matrix is easy and safe, resistant to chemical degradation and the leakage of the dye is easy and recognizable, however it is not possible to predict accurately which protein will bind to these dyes because as we said the binding is highly unspecific and a lot of proteins could be retained by the column. ▪ Metal chelate affinity chromatography: it is based on the principle for which a lot of proteins contain basic groups, most of the times coming from histidine residues. These groups can bind through weak bonds to metal ions. So we can bind to the matrix beads, through the use of a meta chelator, metals like zinc, nickel and copper, and proteins with basic groups will be thus retained in the column. Elution is performed by lowering the buffer pH. This type of chromatography is usually used in the first steps of purification if we know that the protein that we want to isolate has a lot of histidine residues.  ▪ Chromatofocusing: it is based on different pH beads positioned in order in the column, from a more acidic pH to a more basic pH (the matrix is positively charged). The separation is based on the isoelectric point of the protein: since the beads are positioned from a more acid pH to a more basic one, the proteins which have a high isoelectric point will be charged positively when the pH of the column is lower than their isoelectric point, so they will have a positive charge that is repulsed by the matrix; proteins which have a lower isoelectric point will charge negatively at pH values that are higher than their isoelectric point, so they will interact with the positive matrix. So negatively charged proteins absorb to the anion exchanger, positively charged proteins flow down until they reach a point where the pH value equals their own PI. Elution is obtained by changing the pH of the buffer. The advantage is that this technique is high resolving, however proteins precipitate more easily at their isoelectric point and the low salt concentration used in the buffer can induce aggregation (detergent addition can help). Moreover, on an industrial scale this technique is not so convenient, because of economic factors (matrix and eluent are expensive) and complexity factors (the set of the pH gradient is not easy). However, this technique is effective when used in conjunction with other chromatographies. ▪ High performance liquid chromatography (HPLC): it is not a different type of chromatography, it’s just a different way of performing traditional chromatography. So it can be applied to any type of chromatography, the difference is in the elution time because it uses a pump that allows to reach high pressure to elute proteins (enhanced separation in shorter periods of time). It is probably the most widely practiced form of quantitative, analytical chromatography today due to the wide range of molecule types and sizes which can be separated using HPLC or variants of HPLC. For example, insulin and IL-2 are purified with this technique, because it allows us to reach a level of purity that is really close to 100% (the higher is the number of times that the protein has to be administered to the patient, the higher level of purity we need to reach with the downstream process). Various chemical groups may be incorporated into the matrix beads technique such as ion exchange, gel-filtration, affinity, hydrophobic interaction and reverse-phase chromatography. The advantages are that is has a superior resolution due to the reduction in bead particle size, resulting in sharper peaks, the increased flow rates improve fractionation speeds (from hours to minutes) and it has a high degree of automation (good for industrial production scale). The drawbacks are that it is expensive and that it is employed almost exclusively in downstream processing of low-volume, highly-value proteins. Sometimes, the proteins that we have to purify, are not suitable to be isolated from the others as they are (because of their particular chromatographic fingerprint they would require more than 4 chromatographic steps, which are a lot). So, to isolate them we can add a tag: through genetic engineering techniques we induce the incorporation of protein tags on the aminoacidic sequence of the protein of interest, and then we exploit these tags and their characteristics to isolate our recombinant drug. For example, we can add a tag of poly-arginine or poly-lysine, so that we can use their positive charges to perform an ionic-exchange chromatography; or we can add a hydrophobic amino acid tag to exploit it to perform a hydrophobic interaction chromatography; or we can add a poly-histidine tag to the protein so that we can perform the metal chelate chromatography to isolate it. However at the end of this process, our protein of interest will still have the tag attached, and it needs to be removed because it could be immunogenic. How do we remove it? We could use chemical agents (like cyanogen bromide or hydroxylamine) or enzymatic agents (endopeptidases, like trypsin, factor Xa or enterokinase, or exopeptidases, for short tags). Enzymes are more effective than chemical agents but obviously we have to pay attention because enzymes could possibly cleave also our recombinant protein. After the tag is cleaved from the protein we remove it from our final product by performing another chromatography. All of this suggests that the process of production of a biotech drug must always be preceded by a step of study of our protein and of its characteristics, in order to know if we have to use a tag (that must be expressed when we produce the protein in the cell lines, so it has to be prepared during the cloning of the gene). Once our protein is purified after all the chromatographic steps, it will be in solution, in a buffer that needs to be removed. In this process we also have to be sure that the protein is sterile (sterile product means that we have removed all the possible bacteria, viruses or infectious agent that could contaminate our drug). Moreover, we know that proteins can be easily denatured and their activity can be easily altered, with chemical agents (oxidizing agents, detergents), physical agents (extreme levels of pH and elevated temperatures) or biological agents (proteolytic degradation) for example, so we also have to make sure that the protein won’t be damaged and denatured during all of these processes, and we do that by removing the possible damaging agents and avoiding extreme conditions. So the final product formulation involves the addition of various excipients, the filtration of the final product through a 0,22 micrometres absolute filter in order to generate a sterile product, followed by its aseptic filling into final product containers, the lyophilization if the product is to be marketed in a powdered format (how stable is the protein in the solution determines the decision to market the product in liquid or powder form). Before the lyophilization though, we need to stabilize the protein and its function, because different molecular mechanism can underpin the loss of biological activity of any protein. These mechanisms can be divided into two groups: covalent modification and non-covalent modifications. Non-covalent alterations are partial or complete denaturation of the protein; covalent alterations involve hydrolysis, deamidation, imine formation, racemization, oxidation, disulphide exchange, isomerization and photodecomposition. At the end of each chromatographic process we are still not at the highest level of purification that we could reach (especially in the initial steps), so this means that some other proteins, in particular enzymes, could remain in the final solution. If the enzymes are proteolytic enzymes (in particular serine proteases, cysteine proteases or metalloproteases), the protein could undergo proteolytic degradation (especially if it is denatured). This is why every step that follows the chromatographic separation must be performed at low temperatures, so that proteases do not activate. Other precautions that we could use to avoid the activation of these enzymes are for example to minimize the processing times and to use specific protease inhibitors (however some of them are toxic, so inappropriate for the biopharmaceutical processing). Another problem that could occur in the chain of a protein is the deamidation: the aminic residues of asparagine and glutamine can be deamidated (by hydrolysis) and so the two amino acids can become acids (aspartic acid and glutamic acid respectively). This could happen in insulin for example. Oxidation is another covalent alteration that could occur: in the presence of oxygen (caused by air), sulphur atoms present in methionine or cysteine can oxidize, and sometimes disulphide bonds can be formed. Oxidation of methionine is favoured under conditions of low pH, and in the presence of metal ions. For example we could have the oxidation of all three methionine residues in hGH (human growth hormone), resulting in an almost total inactivation of the molecule, while 2 oxidations out of 3 do not affect hGH activity. Oxidation can be best minimized by replacing the air in the headspace of the final product container with an inert gas such as nitrogen. Disulphide bonds can also be formed inside a chain (intrachain), changing the tertiary structure of a protein and resulting, most of the times, in the aggregation of individual molecules, while if