Control of nanopore formation using external triggers
Mutter, Natalie
DOI:
10.33612/diss.131163011
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Publication date:
2020
Link to publication in University of Groningen/UMCG research database
Citation for published version (APA):
Mutter, N. (2020). Control of nanopore formation using external triggers. University of Groningen.
https://doi.org/10.33612/diss.131163011
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1
Introduction-Design
of new targeted cancer
therapeutics
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Cancer is the second leading cause of death worldwide.1 The most common cancer therapies comprise surgery, radiation therapy and chemotherapy. In particular chemotherapy suffers from severe side effects due to a lack of selectivity and off-target toxicity.2,3 Therefore, an emerging field is the development of targeted cancer therapies.4 A new concept is to develop drugs either small molecular drugs, protein-based drugs or conjugates with tailored activity against targets involved in tumorigenesis, proliferation and metastasis.5,6 Advances in genetically engineering and protein engineering as well as the development of directed evolution leaded to a rise in the design of protein-based drugs.7–9 A crucial step to design new protein-based drugs for targeted cancer therapy is to regulate activity of the applied protein. A simple way is to ensure that the protein is only present in its active form at the desired site of action. Therefore, precise control of the delivery and activation by endogenous or external stimuli at the target location is required.
CONTROL BY LOCALIZATION
Evaluation of human cancer cells identified antigens preferentially or exclusively expressed on the surface of cancer cells.10 An example is the epidermal growth factor receptor (EGFR) involved in cancer cell proliferation.11 EGFR is a transmembrane receptor tyrosine kinase stimulated by its ligands epidermal growth factor (EGF) and transforming growth factor α (TNFα) results in intracellular activation of signaling cascades responsible for growth, proliferation and survival.12,13 Overexpression of EGFR is observed in several different cancer types including breast14,15, head and neck16 as well as in lung cancer17. In another example, the folate receptor is overexpressed in for example ovarian18 and breast19 cancer. Other than EGFR and folate, there are more cell surface receptors deregulated in cancer cells which could be suitable as target, examples include the vascular endothelial growth factor receptor (VEGFR)20, the hepatocyte growth factor receptor (HGFR)21 or the transferrin receptor (Tfr)22. Overexpression of those cell surface receptors correlate often with aggressive growing cancers and poor outcome.15,17,23
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Figure 1. Antibodies in immunotoxin preparation. (A) Comparison of antibody variants and their
fragments. Conventional antibodies consist of light and heavy chains. Each chain is divided into the constant domain (blue) and the variable domain (red). Variable domains of both chains fused can be used as antigen-binding fragment (Fab) or single-chain variable domain (scFv). Camelid antibodies lack the light chain. Nanobodies derived from camelid antibodies are the smallest antigen-binding fragment.24,25(B)
Immunotoxin binds via antibody moiety to cell surface receptor and gets internalized. Diphtheria toxin-based immunotoxins rearrange conformation in the endosome and the toxic domain translocates to the cytosol. Immunotoxins based on Pseudomonas exotoxin A are processed in the cell and traffi c via the Golgi and the endoplasmic reticulum to the cytosol. Free toxin in the cytosol inhibits protein synthesis leading to cell death.26
Monoclonal antibodies (mAb) are proteins produced by the immune system to neutralize pathogens. Naturally mAb consist of two identical heavy chains and two identical light chains connected by disulfi de bonds. Each chain has two regions, the constant domain and the variable domain (Figure 1A).27,28 Th e variable domain confer on the mAb the ability to recognize and bind with high selectivity to their targets.28 It is now a therapeutic strategy to generate mAb that recognize tumor antigens, thus blocking their biological function and marking the cells for the body’s immune system.29 An impairment of mAb is their large size resulting in limited tumor penetration30 and their complex composition including posttranslational modifi cations requiring heterologous expression systems31,32 and sophisticated purifi cation processes33. A good alternative are nanobodies (Nbs), which originated from the variable antibody domain of a single-domain camelid antibody (Figure 1A).24,34 Nbs can also bind selectively to specifi c antigens with high affi nity, but in comparison to mAbs, they are smaller and more stable35 leading to more effi cient penetration of tissue36 and lower immunogenicity37. Additionally, Nbs do not require posttranslational modifi cations, therefore can be easily produced in bacteria.38 However, despite their advantages, mAbs and Nbs show limited antitumor activity, although they can be benefi cial as delivery vehicle for cell-killing drugs.6 In contrast, common cancer drugs, which most oft en have potent activity, do not have a strong selectivity toward target cells.5 Th erefore, a new class of drugs has been prepared where cell-targeting elements are conjugated to drugs39,40, toxins41,42,
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cytokines43 or radioactive particles44 to combine their favorable properties. One such class of drugs are immunotoxins, which most often consist of a cell-targeting element, like a mAb, Nb or growth factor chemically or genetically fused to a potent toxin.26 The first immunotoxins were based on plant toxins like ricin45 or bacterial toxins like Pseudomonas exotoxin A46 and diphtheria toxin47. The internalization of the toxin facilitates the inhibition of protein synthesis and results in cell death (Figure 1B). Therefore, the efficiency of immunotoxins depends on sufficient internalization rates of the target receptor.48,49 Otherwise, these are very potent toxins and small concentrations are enough to kill a cell.50 Therefore, it is important that the targeted antigen is merely expressed on diseased cells. Additionally, still other challenges for immunotoxins need to be overcome like limited capacity of solid tumor penetration and immunogenicity.51 Pore-forming toxins could be an alternative to overcome some obstacles of common immunotoxins.
PORE-FORMING TOXINS
Pore-forming toxins (PFTs) are virulence factors produced by bacteria52, sea anemones53 and other organisms54 and are nowadays under investigation for applications in pharmacology. These proteins are able to form transmembrane channels in membranes of target cells. This results in a change of membrane permeability and to an ion imbalance which can lead to cell death.55 PFTs are produced as water-soluble monomers, and bind to specific receptors on the target cell. Membrane binding favors oligomerization and membrane insertion.56 As bacterial virulence factors PFTs are viable targets to design new antibiotics or vaccines necessary to fight antibiotic resistance.57 Since PFTs kill cells efficiently, they become potential candidates for the development of new immunotoxins for targeted cancer therapy.58,59 Currently, the efficiency of immunotoxins face three major challenges, the ability to penetrate solid tumors, immunogenicity and specificity toward cancer cells.42 The main limitation resulting in poor tumor penetration is the molecular size of immunotoxins.60 On the one hand the size of the immunotoxin could be reduced by using mAb fragments, on the other hand the size could be further reduced by using smaller toxins. Conventional toxins such as Pseudomonas exotoxin A or diphtheria toxin ranging from 38 kDa to 58 kDa are larger than PFTs such as Cytolysin A or Fragaceatoxin C.61 An additional advantages could be that PFTs does not require internalization and facilitate excess to cells within a tumor. Therefore, they could be utilized as delivery system by increasing the permeability of target cells for suicidal cancer gene therapy62 or to import impermeable drugs63. However, the toxic moiety of cytosolic-acting as well as for membrane-acting immunotoxins provokes high immunogenicity. Various strategies were developed to reduce immunogenicity including combination therapy with immunosuppressive molecules64, polyethylene glycol modification65, humanization66,67, or removal of B-and T-cell epitopes68–70. The main reason that only a few PFTs where used in the preparation of immunotoxins is there unspecific
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11 toxicity toward most cells.71 Th erefore, it is important to control pore-formation more specifi cally either by incorporating triggers or switches or by reducing the binding affi nity of the native toxin to the membrane. Additionally, the oligomeric nature of PFTs allows to equip monomers with multiple diff erent targeting molecules to increase specifi city. So far, mAbs or antibody fragments targeting immature T lymphocytes58, leukemic cells72,73, breast cancer cells74, lung cancer cells75, colon cancer cells76, or skin cancer cells77 were genetically fused or conjugated to PFTs of diff erent origins for instance from sea anemones59, bacteria78 or humans72. Th e utilized PFTs can be classifi ed based on their structural features. Th e family of α-PFTs form pores by using helices as transmembrane parts whereas β-PFTs insert into the membrane and form β-barrel structures.79,80 An example of a β-PFT that has been intensively studied is α-hemolysin (αHL).81 In contrast, Cytolysin A (ClyA)82 from
Salmonella typhi 83is an α-PFT. ClyA is secreted as 34 kDa water-soluble monomer (Figure 2A) and oligomerization is triggered by contact to detergent or a membrane. Diff erent oligomeric species84–86 are identifi ed, but mainly dodecameric82 pores are observed with a pore diameter of 3.3 nm at the narrowest position (Figure 2C). Pore formation includes remarkable conformational changes and rearrangements of more than half of the residues (Figure 2B).82 Protomers fi rst assemble into dimers and higher oligomeric forms, followed by the assembly into the full pore (Figure 2D).87
Figure 2. Th e ClyA nanopore. (A) Crystal structure of the water-soluble ClyA monomer (PDB entry 1QOY)
(B) Th e ClyA protomer structure with the N-terminal transmembrane helix results from large rearrangements of the monomer. (C) Crystal structure of the fully assembled dodecameric ClyA pore (PDB entry 2WCD). One protomer is highlighted. (D) Schematic representation of the proposed oligomerization mechanism. Binding of the water-soluble monomers via the β-tongue lead to protomer formation and insertion into the membrane. First protomers than smaller oligomers dimerize to form the dodecameric pore.82,84,87
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Another α-PFTs is Fragaceatoxin C (FraC)88 from the sea anemone Actina fragacea. FraC corresponds to the class of actinoporin (cytolysin) protein toxins.89 FraC is expressed as 20 kDa soluble monomer (Figure 3A) and oligomerizes according to binding to the target membrane. A specifi c characteristic of several actinoporins, including FraC, is that the formation of oligomers depends strongly on the sphingomyelin (SM) content of the lipid bilayer.88,90 FraC is composed of a β-sandwich fl anked by two α-helices. Th e oligomerization includes detachment of the N-terminal helix from the β-sheet rich core region and formation of the transmembrane helix (Figure 3B and 3D). Oligomerization results in a funnel-shaped octameric pore with the narrowest diameter of 1.6 nm (Figure 3C).53,88 Lipids do not only act as receptor for FraC on the cell membrane but they also line the pore wall stabilizing the resulting cation-selective channel.88
Figure 3. Th e FraC nanopore. (A) Crystal structure of the water-soluble FraC monomer (PDB entry 3VWI)
(B) FraC protomer results from a jack-knife like motion of the N-terminal helix. Bound lipid molecules are shown in yellow. (C) Crystal structure of the octameric FraC nanopore (PDB entry 4TSY) One protomer is highlighted. (D) Schematic representation of the proposed oligomerization mechanism. FraC monomers bind to the cell membrane in the presence of sphingomyelin. Membrane-bound FraC monomers form multimeric pre-pores which then undergo a conformational change and insert into the lipid bilayer.88,91
Currently, the crystal structure of the soluble and the oligomeric form is solved only for two PFTs, ClyA82,84 and FraC88. Structural knowledge is benefi cial and helps to engineer PFTs for applications not only in pharmacology, but also in biotechnology.92 Notably, when PFTs are reconstituted in artifi cial lipid bilayer they form nanopores. When a potential is applied across the membrane
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13 containing a single nanopore, an ionic current originates. Then the passage of molecules across individual nanopores results in partial blocking of the pore measured as change in ionic current.93 The specific ionic blockade has been used to detect or sequence molecules at the single-molecule level.94 In particular nanopores, like αHL or MspA were engineered for detection and sequencing of nucleic acid.95–97 Larger nanopores such as ClyA were used to sample proteins and enzymatic reactions.98–100 The smaller FraC nanopore can detect proteins and even peptides of a few amino acids in length.101,102
CONTROL BY PROTEOLYSIS
Immunotoxins based on PFTs showed high toxicities toward cancer cells58,76, however they suffered from unspecific killing of non-target cells due to the ability to bind generally to cell membranes.71 In addition to equipping PFTs with targeting molecules, the incorporation of triggers or switches to control pore-formation can add a second level of control and reduce off-target toxicity. Cancer cells overexpress not only cell surface receptors but also proteases.103 Cancer-associated proteases enhance invasion and metastasis by digesting the extracellular matrix and penetrating the basal lamina.104 Proteases overexpressed by cancers are for example furin105,106, cathepsin B107, matrix metalloproteases108 or urokinase plasminogen activator109. In nature, proteases are used to control the activity of proteins. For example, toxins are often expressed as protoxins and activated by proteolytic cleavage at the target site to prevent damage of the host organism or tissue.110 Also some PFTs like aerolysin111, anthrax112 and Vibrio cholerae cytolysin113 are expressed as inactive form and require proteolytic cleavage of redundant peptide sequences at the C-terminus or N-terminus to generate cytolytic active toxins which than form pores into target cells. Therefore, the same approach can be adapted to design prodrugs that are activated in situ by a specific protease. This approach would reduce side-effects and could be used to improve pharmacological properties, such as solubility, stability or clearance from the body.114,115 Immunotoxins with linkers cleaved by cancer-associated proteases have been prepared before resulting in in situ activation.116,117Also one PFT was inactivated by introducing a redundant peptide sequence and could be proteolytically activated by a cancer-associated protease.73,118,119 Apart from proteases, also other triggers could be suitable to be implemented into drug design to increase selectivity. Other endogenous triggers could be the pH or redox-potential at the target site.120,121 One hallmark of cancers is the dysregulation of the pH, in this regard the pH in the extracellular microenvironment of a tumor is reduced from normally 7.4 to 6.5. The lowered pH can be used as trigger to activate prodrugs or to cause release of a drugs from pH sensitive nanocarriers.122 Additionally, external triggers such as light could add further control.123
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CONTROL BY LIGHT
One possibility to activate drugs selectively and to reduce side eff ects would be to control the drug activity using an external trigger like light. Photopharmacology uses photoswitches to establish control over the action of bioactive molecules.124–126 Photocontrolling the activity of biomolecules
in vivo opens up new opportunities in designing drugs with reduced non-selective toxicity and
to study physiological processes in living organisms. In this regard is light a particular attractive trigger because it is noninvasive, allows bioorthogonal control as well as high spatiotemporal control.124,125,127 Two classes of molecular photoswitches are commonly used either light-responsive small molecules126 or naturally occurring photoresponsive proteins128.
Chemical-based photoswitches
In photopharmacology light-responsive small molecules can be either incorporated into a drug or covalently attached to a biological target.124–126 Several diff erent photoswitches are known like spiropyranes129, diarylethenes130, hemithioindigos131 or fulgides132.
Figure 4. Common photoswitches used in photopharmacology. Chemical structure of an azobenzene
photoswitch. Azobenzenes are thermally stable in the trans state, irradiation with light of a specifi c wavelength results in the metastable cis state.
Azobenzenes are the most commonly used photoswitches, due to their favorable photophysical properties, such as high photostationary states and quantum yields as well as fast photoisomerization and low rates of photobleaching.126,133 Azobenzenes switch between two isomeric forms, the thermally stable trans state, and irradiation with light interconverts it into the metastable cis state. Either thermal relaxation or light irradiation causes the reversion from
cis to trans (Figure 4).133 For an optimal function as a drug, the switch should be non-active in its thermally stable state, while being activated upon irradiation with light.
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Figure 5. Light-controlled pores. (A) An azobenzene blocks a potassium channel more effi ciently as linear
trans isomer than as cis isomer.134 (B) An azobenzene tethered to a potassium channel blocks the ion fl ow
in the trans confi guration, but aft er photoisomerization it is too short to block the channel eff ectively.135 (C)
Attachment of a photocaging group blocks the OmpG pore. Irradiation with UV light results in removal of the photocaging group and in an ion fl ow.136 (D) An agonist is bound to the ligand-gated ionotropic glutamate
receptor via a linker including an azobenzene. While in trans isoform the ligand cannot reach the binding site, photoisomerization results in binding of the agonist and opening of the ionotropic glutamate receptor.137
(E) Th e MscL channel was equipped with a spiropyrane switch. Upon irradiation ring opening takes place resulting in the zwitterionic merocyanine structure and in opening of the MscL channel.138 (F) α-HL pore
forming abilities were inactivated by attachment of a photocaging group to the glycine-rich loop. Uncaging of α-HL by irradiation results in pore formation.139
Most photoswitches are introduced into the structure of known small molecular drugs.140–142 Th e approach has been successfully demonstrated in developing light-controlled anticancer drugs.143–146 Latterly, photoswitches are covalently attached to proteins to successfully stabilize their structure147 or to modulate their function148. In particular, photoswitches have been attached to PFTs such as ion channels, which are interesting pharmaceutically targets because they play important roles in cellular communication149,150, exchange of material151 and as defense agents56. Incorporation of photoswitches allowed controlling the function of channels125 including the light-dependent blocking of the ion fl ow through potassium channels134,135,152,153 (Figure 5A and 5B), voltage-gated proton channels154, P2X receptors155, Ca2+ channels156or OmpG136 (Figure 5C). Th e opening and closing of the ligand-gated ionotropic glutamate receptor137,157–159 (Figure 5D) or
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the mechanosensitive channel MscL138,160 (Figure 5E) can be controlled by light aft er attachment of photoswitches to specifi c positions within the proteins. Additionally, chemical photoswitches were used to induce light-controlled protein translocation by the SecYEG complex161 or to design a light controlled pH valve162. Another attempt, which might be particularly attractive in photopharmacology, was to control the formation of a PFT using light irradiation. It was shown that pore-formation of α-HL can be prevented by attaching a photocleavable 2-nitrobenzyl group to a glycine-rich central loop which lines a segment of the transmembrane pore. Subsequent light irradiation cleaved off the protective group and activated α-HL resulting in hemolytic activity (Figure 5F).139 However, pore formation could not be controlled in reversible manner, which is necessary to design highly selective cancer drugs with low off -target toxicity. Currently, the application of most chemical photoswitches in vivo is limited due to UV light depended photoisomerization, which can be toxic to healthy cells163 and has a limited penetration depth164.
Protein-based photoswitches
Figure 6. Th e AsLOV2 domain. (A) Simplifi ed AsLOV2 photocycle. In the dark the oxidized FMN is
noncovalently bound to the protein. Blue light irradiation results in reduction of the FMN due to covalent bond formation between a conserved cysteine residue and the C4a of the FMN. (B) Schematic structure of the AsLOV2 in the dark (left ) and lit state (right). In the dark, the FMN (red) in noncovalently bound. Blue light irradiation results in bond formation between the FMN and a cysteine residue (green sticks) and induces unfolding and displacement of the C-terminal Jα-helix (dark blue). (PDB: 2V0U)169,170
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17 Photoresponsive proteins may offer an alternative to control the activity of biomolecules with light. Photoreceptor proteins are found in plants, algae and bacteria fundamental for photosynthesis and photomorphology.165 These proteins utilize a cofactor, which is required to sense light.128 Light induces changes of the photosensory domain resulting in structural metamorphosis of the functional domain and a change in activity. Several different photosensory domains evolved in nature basing on different chromophores. Phytochromes166 absorb red light via a tetrapyrrole cofactor, while cryptochromes167 and phototropins168 are activated by blue light absorbed by a flavin chromophore.
A heavily studied photoactivatable protein is the photosensory domain of phototropins, the light-oxygen-voltage domain (LOV). LOV sensors are a subclass of the versatile Per-ARNT-Sim family171 and control a variety of different effector domains ranging from kinases, phosphodiesterases, sulfate transporters to DNA-binding proteins.172 Typically LOV domains contain the flavin binding consensus sequence GXNCRFLQ, which includes a cysteine that is important for the photocycle and in signal transduction.173 Light absorption of the chromophore results in bond formation between the flavin and the conserved cysteine residue of the LOV domain. Flavin-thiol adduct formation induces major structural rearrangements, including unfolding of helices169 or dimerization174. In particular, the LOV2 domain of Avena sativa (AsLOV2) has been extensively studied and used to design new photoswitchable proteins. Upon blue-light irradiation the flavin mononucleotide gets excited and binds via the C4a of the isoalloxazine ring to the cysteine of the AsLOV2 domain.175 This promotes unfolding and detachment of the C-terminal Jα-helix from the β-sheet core.169 The light-activated state is metastable and in the dark it thermally decays to the ground state.176 The AsLOV2 domain was genetically fused to a number of proteins to photoregulate their function. One of the first examples linked the AsLOV2 domain to the E.
coli trp repressor to build the light-regulated DNA binding protein LOV-TAP (Figure 7A).177 The two proteins were fused such that they share a mutual α-helix, while in the dark the shared helix contacted the AsLOV2 domain resulting in an inactive trp repressor. Blue light irradiation unfolded the Jα-helix, allowing correct folding of the trp repressor and DNA binding. Furthermore, the catalytic activity of metabolic enzymes, such as dihydrofolate reductase178 and pyruvate kinase179 could be allosterically controlled, therefore the AsLOV2 domain was inserting into a surface loop sensitive to modifications. Irradiation with blue light enhanced the activity of the chimeric enzymes, however the enzymes showed reduced substrate affinities.
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Figure 7. Optogenetic approaches to control protein activity. (A) Th e LOV-TAP system can control
transcription. Th erefore, the AsLOV2 domain was fused to the trp repressor sharing an α-helix. Upon irradiation the Jα-helix of the AsLOV2 domain unfolds and allows correctly folding of the repressor and binding to DNA.177(B) Optogenetic control of the membrane potential was achieved by fusing the C-terminus
of the AsLOV2 domain with the N-terminus of a potassium channel. Irradiation with blue light results in opening of the channel and increased potassium conductance.180 (C) Th e GTPase Rac1 was fused to a AsLOV2
domain. Th e AsLOV2 domain sterically blocks all interactions until irradiation unwinds the Jα-helix. Rac1 regulates for example cell migration and other processes.181 (D) Intracellular calcium concentration was
controlled by fusing calmodulin to the AsLOV2 domain. In the dark state the Ca2+ is bound to calmodulin,
light-irradiation resulted in the destabilization of calmodulin and release of Ca2+.182 (E) Th e opening of
Ca2+- specifi c channels in the plasma membrane was induced by binding to photoregulated STIM peptide.
In the dark the STIM peptide was shielded by the AsLOV2 domain. Upon irradiation with blue light the STIM peptide becomes accessible for interactions with the Ca2+- specifi c channel.183 (F) Th e localization
of proteins inside a cell was controlled using the TULIP system. In the dark the LOVpep is caged upon irradiation the Jα-helix undocks and exposes the epitope for binding ePDZ, which recruits the protein of interest.184 (G) Th e LOVTRAP system controls the localization of proteins inside a cell. In the dark the protein
of interest is bound to the AsLOV2 domain via ZdK. Irradiation with blue light results in dissociation and enable the protein of interest to reach the target site.185 (H) A protein of interest was fused to the AsLOV2
domain equipped with a degradation signal. In the dark the degradation sequence is buried, upon blue light irradiation the sequence can be recognized and the protein gets degraded by the proteasome.186
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19 In optogenetics AsLOV2 fusion proteins are used to achieve optical control over cellular behavior. Different approaches have been employed to control biological processes like neuronal activity.180,187 Typically, the light-sensitive module is directly fused to a protein of interest.181,188 Thus, a light-gated potassium channel was prepared by fusing the C-terminus of an membrane tethered AsLOV2 domain with the N-terminus of a potassium channel.180 Irradiation with blue light resulted in increased potassium conductance (Figure 7B). Furthermore, the activity of the small GTPase Rac1, important for regulating cytoskeletal dynamics, was controlled by light. The GTPase Rac1 was fused to the AsLOV2 domain, which was sterically blocking Rac1 interactions until irradiation unwound the Jα-helix linking AsLOV2 to Rac1 (Figure 7C).181 Besides that the accessibility of regulator molecules, such as calcium ions (Ca2+) can be controlled.182,183 On the one hand, the AsLOV2 domain was inserted into the Ca2+-binding calmodulin. In the dark calmodulin bound the Ca2+, upon photoirradiation the structural changes in the AsLOV2-calmodulin fusion protein caused the release of Ca2+ (Figure 7D).182 On the other hand, the opening of Ca2+-specific channels was gated via interactions with the stromal interaction molecule (STIM) fragment. The Ca2+ sensing STIM peptide was shielded by fusion to the Jα-helix of the AsLOV2 domain. Blue light irradiation resulted in exposure of the STIM peptide, binding to the Ca2+-specificchannel and influx of Ca2+ ions (Figure 7E).183
Alternatively, the localization of proteins in the cell can be controlled using recruitment methods like tunable, light-controlled interacting protein tags (TULIPs)184, improved light inducible dimers (iLID)189 or LOV2 trap and release of proteins (LOVTRAP)185. In the TULIPs system a peptide was caged by fusion to the Jα-helix of the AsLOV2 domain (LOVpep), upon photoexcitation the peptide got unmasked and bound to ePDZ, a protein domain interacting specifically with peptides. Proteins fused to ePDZ could get recruited by blue light to target locations where LOVpep was tethered (Figure 7F).184 The iLID system works similar utilizing a different peptide and corresponding binding partner.189 In contrast, the LOVTRAP system used the small protein ZdK, which selectively binds the dark state of the AsLOV2 domain. The protein of interest was fused to ZdK and sequestered by binding to a membrane tethered AsLOV2 domain, upon irradiation ZdK dissociates and the protein of interest could move to its side of action (Figure 7G).185 Additionally, the intracellular concentration of a biomolecule of interest can be regulated by light-sensitive transcription177,190, translation191 or degradation186. Therefore, the AsLOV2 domains was fused to transcription factors177 (Figure 7A) or translation repressor proteins191 to inactivate their function. Light induced structural changes of the AsLOV2 domain resulted in activation of the chimeric proteins. Alternatively, the protein of interest was fused to the N-terminus of the AsLOV2 domain, which in the dark buried a degradation signal. Irradiation results in exposer of the degradation signal and in proteasomal degradation of the protein (Figure
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7H).186 Latterly, photoreceptors are also employed to design selective drug delivery systems or new photoresponsive protein-based drugs to reduce off-target toxicity.192
In optogenetics AsLOV2 domain and other photoreceptors have versatile applications.193 The next step will be to introduce multiplexing, to investigate or control several biomolecules at the same time, to gain insights into complex biological sytems.194 Therefore, new photosensory domains, and photoreceptor systems with high wavelength selectivity and low dark state activity need to be investigated.
DIRECTED EVOLUTION
The development of protein-based therapeutics as important class of medicine was facilitated by advances in molecular biology techniques, due to the high demand to optimize multiple characteristics of protein therapeutics.7–9 A powerful method to improve or alter the activity of proteins for industrial195 and therapeutic applications196 is directed evolution. It mimics the process of natural selection to obtain proteins with user-defined functions. Directed evolution has been successfully used to improve for example protein stability197,198, or binding affinities199,200 and to alter substrate specificities of enzymes201,202. Additionally, it was utilized to tailor PFT properties. For example, the solubility and electrical properties in lipid bilayers of ClyA86 were improved, or the hemolytic activity of αHL was increased203. Additionally, a tumor-protease-activated αHL was prepared119 and anthrax toxin was evolved with increased selectivity toward targeted cancer cells204. Directed evolution is advantageous for a large number of applications including the engineering of optogenetic tools. It was used to improve properties such as the brightness205,206, photostability207, thermostability208 and kinetics209 of LOV domains.
Directed evolution improves the protein properties in iterative cycles consisting of four steps (Figure 8). In the first step genetic diversity is generated by mutagenesis. Then protein variants from a library of mutated genes are expressed in suitable host organisms. In the third step, high-throughput screening or selection of proteins for a desired property is applied in order to discover improved variants.210 Preferably, the chosen protein of interest has already a low level of activity for the desired reaction at the begining.211 In the last step, the genes encoding for improved variants are selected, and used for another cycle of directed evolution.210
DNA-library construction methods
Complete randomization of a 100 amino acid long protein would yield 20100 possible different variants. Therefore, it is impossible to test the entire mutational space of a protein, because it would
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21 exceed the feasible library size. Instead, improved proteins result from stepwise introduction of several benefi cial mutations which accumulate over many generations.210 Th e occurrence rate of spontaneous mutations is very low, therefore diff erent techniques are necessary to increase genetic diversity to generate libraries.
Figure 8. Directed evolution. Th e workfl ow of directed evolution experiments consisting of four iterative
steps. First genetic diversity is generated typically by random mutagenesis. Th en the gene library is expressed in a suitable host organism. Next comes one of the key steps, which is the screening or selection of functional variants. Th e genes encoding for the selected variants are then replicated and serve as template for the next cycle. It is crucial during directed evolution to maintain the consensus between genotype and phenotype.210
Random mutagenesis is the method of choice in the absence of knowledge about the structure-function relationship. Th e most commonly used random mutagenesis strategy is error-prone polymerase chain reaction (PCR).212,213 Point mutations can be generated during PCR amplifi cation using a low fi delity DNA polymerases like Taq214 and by changing the PCR conditions to reduce the fi delity of the DNA polymerase. For example, increased mutation rates can be caused by increased magnesium concentrations215, addition of manganese216 or the use of mutagenic dNTP analogues217. Mutations accumulate with each PCR cycle, therefore increasing the number of PCR cycles results in an increased average number of mutations per gene.
Alternatively, mutations can be targeted to a specifi c position. Th is can be useful if residues important for protein function are known, or if the protein structure is solved and such residues can be identifi ed. Th en, focused mutagenesis toward such residues allows sampling of the
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full amino acid repertoire and can provide a better chance to improve variants. The strategy of choice is to perform site-directed saturation mutagenesis using primers containing one or more degenerated codons.218–220 This approach allows the introduction of simultaneous targeted modifications at multiple residues, which can result in combinations of mutations with synergistic effect.221 Computational tools can facilitate the design of focused mutagenesis libraries. It can be employed to predict key residues in the protein responsible for binding, activity or stability.222–224 Thus, the library size can be reduced and variants with improved properties enriched.
Another method used in directed evolution experiments is DNA shuffling, which was developed by Stemmer225,226 in 1994. It mimics the natural process of homologous recombination to result in new beneficial combinations of mutations. This approach requires the fragmentation of a set of homologous genes by DNase I and random reassembly by PCR resulting in chimeric genes. Since then several other DNA shuffling methods using different fragmentation or reassembly techniques were developed. In random chimeragenesis on transient templates (RACHITT)227 DNA is fragmented by DNase I and hybridized on a single-stranded uracil-containing scaffold. Then overhangs are trimmed, the gaps are filled, the fragments ligated and finally the scaffold digested. Thus, the number of crossovers could be increased and the number of unshuffled genes reduced. Staggered extension process (SteP)228 is a recombination method performed in a single tube, based on a modified PCR protocol. The extension step is interrupted prematurely by heat denaturation then fragments can anneal to a different template and get further extended. This process is repeated until full-length chimeric genes are produced. The above-mentioned recombination techniques are only effective, when a diverse population of starting genes is used. Either naturally related homologue sequences or sequences derived from previous libraries can be employed as starting genes.229,230 In contrast, also homologous recombination can be performed using non-homologous random recombination (NRR).231,232 Sequences are fragmented by DNase I followed by sequence-independent blunt-end ligation to generate hybrid genes. NNR allows rearrangement of sequence motifs, insertions or deletions, but a majority of non-homologous recombinants will be disrupted and unable to fold.
Latterly, researchers increasingly use targeted mutagenesis, due to the increasing number of available structures and advances in computational modeling. However, deducing protein function from the amino acid level is challenging. Besides, random mutagenesis can probe mutations distant from functional sites. Therefore, the most effective strategies for library design combine knowledge-based and random mutagenesis.233
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23 Selection and Screening methods
Successful directed evolution experiments depend mainly on high-throughput selection or screening of the prepared library to identify improved variants. Nobel prize winner Frances Arnold phrased ones “you get what you screen for” 234 and nowadays this is considered as the first law of directed evolution. Consistently, methods of library analysis need to be developed and adapted for every protein of interest. An important feature necessary during laboratory evolution is to maintain the genotype-phenotype association.235 One way to achieve this is using spatial separation of the individual mutants. Variants are expressed in an unicellular organism and each mutant is screened as colony on solid media119,197,236,237 or in multiwell culture plates86,238,239. The advantage of using multiwell culture plates is the compatibility with many different assay techniques, like spectroscopic239,240 and fluorescent readouts209,239, nuclear magnetic resonance (NMR)241, high-performance liquid chromatograph (HPLC)242, gas chromatography243,244 or mass spectroscopy245. However, although almost any protein activity can now be screened, the size of the library is usually limited to ~104 mutants per screening round.210,246 Higher throughput can be accomplished using assessment of optical features like color247, fluorescence229, luminescence248 or turbidity249. Either the protein of interest has naturally an observable phenotype, like GFP250, rhodopsin251, or subtilisin247, or a detectable reporter needs to be used whose signal is proportional to protein activity252–256. PFTs function can be easily evaluated by hemolytic activity86,119, following the release of hemoglobin from red blood cells by measuring the change in the solution absorbance.257 Alternatively screening by flow cytometry258 avoid the spatial separation of the clones but requires fluorescent reporters to identify and isolate cells containing the desired variants259,260. Flow cytometry is a powerful tool in combination with yeast surface display, where the protein of interest is displayed as fusion to the cell surface protein Aga2 of yeast.261 This method had enabled the discovery of new protein-protein interactions262, the development of antibody-antigen pairs199,263 and latterly the evolution of bond-forming enzymes264,265 or proteases201. Likewise, flow cytometry can be combined with in vitro compartmentalization266 to facilitate the use of fluorogenic substrates. Therefore, genes, the transcription machinery and substrates are entrapped in either a water-oil-water droplet253 or in polyelectrolyte shells267.
While screening requires evaluation of every protein for the desired property, selection eliminates nonfunctional variants automatically.268 As for screening methods also selection methods require coupling of phenotype and genotype. In selection methods, such as cell surface display261,269 or phage display200,270 a cell or bacteriophage serves as compartment to connect gene and protein. For example, in phage display the protein of interest is expressed on the surface of the bacteriophage coat through fusion to coat protein III.270 A common application is to select for binding affinities using affinity purification (biopanning).271 Therefore, the target is immobilized, low affinity
1
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binders are washed away, while high affinity binders are eluted and used to infect E. coli to produce an enriched library. The cycle is repeated several times to select for high affinity binding peptides or proteins. Phage display is one of the most commonly used display method in particular to develop new therapeutic antibodies.200,272,273 Nowadays several display methods suitable for directed evolution experiments are developed like bacterial display274, ribosome display275 or mRNA display276.
Another selection method is to couple the activity of the protein of interest to the survival of the organism. It is easily feasible for enzymes neutralizing or exporting antibiotics225,277 as well as for metabolic enzymes278. However, it is more challenging to design selections for other protein activities, one option is to link the desired activity to the expression of an antibiotic resistance gene.279,280 Limitations of in vivo selection, such as limited transformation efficiencies or host genome mutations, can be bypassed by in vitro approaches. Library members are translated in artificial compartments and activity is coupled to PCR amplification of the corresponding gene. Enhanced activity of a library member results in enrichment of their encoding gene.266,281 Therefore, this methods is utile for DNA acting enzymes282,283 or can be used to evolve enzymatic reactions which control the expression of polymerases281,284.
Directed evolution is an amazing tool to optimize protein properties or functions and to create proteins with new functions. However, for each protein of interest the optimal screening and selection technique, as well as the suitable genetic diversification strategy needs to be chosen in accordance with the protein features.
SCOPE OF THIS THESIS
In this thesis we describe a method to design new protein-based therapeutics on the basis of pore-forming toxins. In a modular approach the pore-forming toxin was equipped with different targeting components and directed evolution was used to fine -tune protein properties.
Chapter 1 provides a review on the current state of controlling pore-forming protein activity to design new targeted cancer drugs. Focusing on three approaches to increase selectivity of drugs. First by controlling localization of the drug. Second by selective activation by proteases at the target site. Third by controlling activity using the external stimulus light. Additionally, the possibilities to fine-tune protein-based drugs by directed evolution are depicted.
Natalie_Proefschrift.indd 24
25 Chapter 2 demonstrates the synthesis and characterization of a new class of immunotoxins comprising of a pore-forming toxin. Therefore, PFTs were either covalently attached to folate or genetically fused to a nanobody against EGFR to target specifically cancer cells. Conjugation resulted in a two-fold increase in cytotoxicity toward cells expressing the cognate receptors. Additionally, in a second approach we suppressed off-target activity, therefore the toxin was synthesized as protoxin. We introduced a polypeptide segment at the transmembrane N-terminus to cap activity. In situ proteolysis by the cancer-associated protease furin resulted in activation of the toxin. Further directed evolution was used to fine tune immunotoxin properties, to improve soluble expression, increase target affinity and to reduce off-target activity.
Chapter 3 describes the preparation of a photocontrolled nanopore. The pore-forming toxin FraC was equipped with photoswitchable azobenzenes to remotely control nanopore assembly by light. Therefore, we attached photoswitchable azobenzene pendants covalently to cysteines introduced at various positions near the sphingomyelin binding site of FraC. Cytolytic activity of FraC attached to the azobenzenes was tested using hemolytic assays. Several constructs showed the desired properties, as they were less active in the thermal resting state (trans isomer) than in the light excited state (cis isomer). Notably, the para-sulfonated azobenzene attached to position Y138C of FraC showed a five-fold higher hemolytic activity in cis isoform compared to trans isoform. Additionally, cytotoxicity of photocontrolled FraC could be stopped by irradiation with white light, hence providing reversible control of nanopore formation.
Chapter 4 presents the design of a genetically encoded pore-forming toxin activated by both proteases and light of a variety of wavelengths. A photoresponsive AsLOV2 domain was genetically fused to the N-terminus of FraC, which suppressed the cytotoxicity of the protein fusion. Irradiation with blue light results in conformational changes of the AsLOV2 domain and the exposure of a protease cleavage site. Simultaneous exposure to a cancer protease and light induced in situ proteolytic cleavage activating FraC and causing cellular cytotoxicity. Interestingly, random mutagenesis coupled to DNA shuffling and subsequent screening on red blood cells allowed identifying variants activated by green and even red light. New green light responding variants were successfully employed to target specifically cancer cells.
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