ABSTRACT
3. Proteins Assemble into Dynamic Nanomachines
First, self-association of monomers occurs through hydrophobic domains in a process known as coacervation. Second, lysine residues of elastin are cross-linked by one or more members of the lysyl oxidase family. The non- elastin component of elastic fi bers is collectively referred to as microfi brils and includes fi brillins, fi bulins, and microfi bril-associated glycoproteins.
The peripheral microfi brils around elastin in elastic fi bers may contribute to elastin fi ber assembly as a guiding scaffold, as the microfi bril appears prior to elastin deposition. The elastin can elastically extend and contract in repetitive motion when hydrated. It has been suggested that elastin functions as an entropic spring. Upon relaxation from a stretching, elastic recoil is driven by increasing entropy that complies with increased disorder of the polypeptide chain and of surrounding water molecules. This implies certain degree of structural heterogeneity and disorder within the elastin monomer, where the formation of extended secondary structures is restricted in favor of transient and fl uctuating local motifs.
Water-swollen elastin fi bers are about 5–8 mm in diameter. Negative staining electron microscopy of elastin fi bers revealed distinct fi laments aligned parallel to the long axis of the fi ber (Gotte et al. 1974, Gotte et al.
1976). Similar fi laments were found in α-elastin, coacervates of tropoelastin, synthetic elastin-like polypeptides, fibrous elastin, and tropoelastin (Muiznieks et al. 2010 and references therein). These fi laments laterally associate into fi brils, which are bridged at regular intervals, and display a high propensity to aggregate into thick bundles (Pepe et al. 2007). Stretching of elastin fi bers reveals a transition from a glassy, plastic, disordered network to well-ordered fi laments aligned along the direction of the applied force.
The solution structure of individual hydrophobic domains of tropoelastin and elastin-derived peptides is disordered and fl exible in conformation. The fl uctuating β-turns within hydrophobic elastin-like sequences are believed to be labile and may contribute to elasticity. The elastin monomer is fl exible and highly dynamic. Structural disorder within monomer and aggregated elastin does not preclude the formation of preferred self-interactions or specifi c interactions with matrix and cell components, and thus the directional formation of fi bers. A high degree of combined composition of proline and glycine residues prevents the collapse of hydrophobic residues into compact globular structures by limiting the formation of backbone self- associations. Thus, the elastin monomer remains more hydrated and fl exible than globular proteins of similar size, where its conformational disorder plays an essential role in driving extensibility and elasticity.
homeostasis. Most natural biological molecular nanomachines are enzyme proteins. Enzymes catalyze biochemical reactions in cells, converting substrates into products. Almost all metabolic processes in cells are accomplished by enzyme proteins. Binding substrate often induces conformational transition in an enzyme protein, which lowers the activation energy of the biochemical reaction. The substrate-bound conformation of an enzyme may form an electrostatic complementarity to the transition state of the substrate, or form a covalent intermediate to provide a lower energy transition state, or mechanically distort the substrate into their transition state. Enzymes often exhibit complex internal dynamics required by their functionality, a common property shared by all types of protein nanomachines.
All eukaryotic cells contain cyclic molecular motors, a special class of enzyme proteins that convert chemical energy into mechanical energy or vice versa. Different from structural proteins that assemble into relatively static high-order architecture, many protein nanomachines are highly dynamic and can generate forces. They may interact with high-order protein architecture to fulfi ll their cellular functions and perform mechanical work during enzymatic cycle. For instance, a golf-club shaped protein myosin uses actin fi laments as tracks to crawl bipedally. Myosin, kinesin and polymerases are the remarkable examples of cyclic motors. They can take an infi nite number of cyclic steps of conformational changes without damaging the integrity of their structures, given suffi cient quantities of
“fuel” molecules, such as adenosine triphosphate (ATP). Therefore, the natural protein nanomachines have been considered a primary source of nanoscale engineering of molecular machinery and studied extensively from the standpoint of nanomaterial engineering.
3.1 Nanochannels and Nanopumps
There are three principal classes of membrane transport proteins: channels, transporters, and pumps (Hille 2001). These proteins reside in the plasma membrane and in the membranes of intracellular organelles, lysosomes, and mitochondria. They are extremely important for a myriad of cellular functions, ranging from uptake of nutrients such as glucose to more complex physiological tasks such as the reabsorption of solutes by the kidney and the propagation of action potentials in neural cells. Ion channels catalyze the rapid, selective and passive transport of ions down their electrochemical gradients (Jentsch et al. 2004). They vary in their degree of selectivity: some are highly selective for K+, Na+, Ca2+, Cl– ions, or H2O, whereas others are selective for certain anions or cations. Selectivity is enforced by a region in ion channels called the selectivity fi lter, which makes partial dehydration of the permeating ions energetically favorable over other ions of similar
size. Ion channels undergo regulated opening and closing in a process called gating. They may be gated by ligand, voltage, mechanical stretch, or temperature change. The general architecture for several ion channel proteins have been characterized, and their atomic models have been built to account for their selectivity, transport energetics and gating mechanism.
Potassium (K+) channels are the most widely distributed type of ion channels and found in nearly all living organisms. K+ channels are found in most cell types and control a wide variety of cell functions.
They form K+-selective pores across cell membranes. Each K+ channel is a homotetramer, with the four subunits together forming a central pore, through which ions pass (Doyle et al. 1998, Zhou et al. 2001). Each subunit has two transmembrane α-helices separated by a pore (P) loop. The P-loop contains a short helical element called the pore helix. The pore consists of two major components, the selectivity fi lter and the central cavity.
The selectivity fi lter, which is the narrowest part of the pore and is 12 Å long and 3 Å wide, is near the extracellular opening and binds K+ ions (Fig. 3a). Six ion-containing sites have been observed in the channel: four
Figure 3. Nanochannels and nanopumps. (a) A crystal structure of the potassium channel from Streptomyces lividans, with sequence similarity to all known K+ channels, particularly in the pore region (PDB ID: 1BL8) (Doyle et al. 1998). (b) A crystal structure of the Calcium ATPase pump from skeletal muscle sarcoplasmic reticulum (1IWO) (Toyoshima and Nomura 2002).
(c) A crystal structure of α-hemolysin, a heptameric transmembrane pore (PDB ID: 7AHL) (Song et al. 1996, Mathé et al. 2005), which has been extensively used in nanopore devices for single-molecule analysis and biosensing (Clarke et al. 2009, Wang et al. 2011, Rodriguez- Larrea and Bayley 2013). In all panels, the dashed lines approximately mark the position of lipid bilayer membrane.
sites (P1–P4) within the selectivity fi lter, one on the extracellular side of the fi lter (P0) and one in the central cavity (P5). These sites represent the transport pathway of K+ ions. K+ ions in solution are hydrated. Dehydration of ions is energetically disfavored because it must remove the partial negative charges of water dipoles from positively charged ions. The partial negative charge of the oxygen atoms in the selectivity fi lter imitates water molecules, creating a hydrophilic environment that lowers the dehydration energy for the permeating potassium ions and mimics the weak negative charges of water dipoles forming the hydration shell (Berneche and Roux 2001).
K+ channels open and close within milliseconds to maintain the negative resting membrane potential and prevents ion leak. Different subfamilies of K+ channels are gated by different extracellular or intracellular signals. For some K+ channels, the gate opens intrinsically with a sensing mechanism that links channel activity to the metabolic state of the cell. In another type of gating mechanism, ligand binding to the channel’s intracellular domains causes a conformational change that opens the channel gate. Ca2+, ATP, trimeric G proteins, and polyamines are examples of such ligands.
In voltag e-gated K+ channels, changes in the membrane potential lead to conformational changes in the transmembrane segments that open the channel gate. An example of voltage gating is the voltage-sensing mechanism of voltage-dependent K+ channels, which maintains the resting membrane potential at negative voltages and allows for the termination of action potentials in electrically excitable cells such as neurons and muscle cells (Jiang et al. 2003, Long et al. 2005a, Long et al. 2005b) . Some K+channels are gated by both voltage change and ligand binding. These gating mechanisms of K+ channels were also found similarly in other ion channels, such as Na+ channels and Ca2+ channels, although others exhibited different structural change to close the channels (Stuhmer et al. 1989, Toyoshima et al.
2000, Toyoshima and Nomura 2002). For instance, voltage-dependent Na+ channels are closed by specifi c hydrophobic residues that block the pore.
In contrast to ion channels that transport ions passively, transporters and pumps are solute-selective carrier proteins that use free energy to actively transport solutes against their electrochemical gradients. These transport proteins often alternate between two different conformations.
In one conformation, the transport protein binds solute molecules on one side of the membrane; and in another conformation, it releases solutes on the other side. Uniporters moves solutes down their transmembrane concentration gradients, such as glucose transporters (Mueckler et al.
1985). Symporters and antiporters move a solute against its transmembrane concentration gradient; this movement is energetically coupled to the movement of a second solute down its transmembrane concentration gradient. Many transporters are part of the major facilitator superfamily (MFS) transport proteins that translocate sugars, sugar-phosphates,
drugs, neurotransmitters, nucleosides, amino acids, peptides, and other solute across membranes. A typical MFS member, the bacterial lactose permease LacY functions as a monomeric oligosaccharide/H+ symporter, whi ch us es the free e nergy released from the translocation of H+ down its electrochemical gradient to drive the accumulation of nutrients such as lactose against its concentration gradient.
Transporters and pumps use different sources of energy to accomplish transport. Transporters couple transport with the energy stored in electrochemical gradients across the membrane. In contrast, pumps use energy from ATP or environmental trigger such as light to drive transport.
A typical molecular pump is Ca2+-ATPase (Fig. 3b), which pumps Ca2+ into intracellular storage compartments using the free energy released by ATP hydrolysis (Toyoshima et al. 2000, Toyoshima and Nomura 2002). In each pumping cycle, the Ca2+-ATPase changes its conformation between two states: the E1 conformation, which binds Ca2+ on the cytosolic side with high affi nity, and the E2 conformation, which binds Ca2+ with a signifi cantly reduced affi nity and therefore releases Ca2+ from the Ca2+-binding sites into the sarcoendoplasmic reticulum. Some molecular pumps are also rotary molecular motors. For example, H+-ATPase pumps protons out of cytosol across the membrane into the organelle lumen through rotational cycle driven by ATP hydrolysis (Wilkens et al. 1999).
In addition to these ion channels, transporters and pumps, a pore- forming β-barrel toxin family has been involved in bionanotechnology development. The fi rst identifi ed member of this toxin family, a heptameric transmembrane protein nanopore α-hemolysin (Fig. 3c) consists mostly of β-sheets with only about 10% α-helices contents (Song et al. 1996, Mathé et al. 2005). α-hemolysin has been used in lab-on-a-chip devices for single-molecule analysis and biosensing, translating sequences of nucleotides or peptides directly into electronic signatures (Clarke et al.
2009, Wang et al. 2011, Rodriguez-Larrea and Bayley 2013).
3.2 R otary Motors
ATP synthase is a rotary molecular motor found on the thylakoid and inner mitochondrial membrane in eukaryotic cells, which couples the free energy of the electrochemical proton gradient across the membrane to ATP synthesis (Noji et al. 1997, Itoh et al. 2004, Stock et al. 1999, Rondelez et al.
2005). It is an extraordinary enzyme that provides energy source for the cells to consume through the synthesis of ATP. ATP is the most commonly used biological “fuel” in cells from most organisms. The ATP sy nthase join adenosine diphosphate (ADP) and phosphate (Pi) covalently to produce ATP. Energy is then released in the form of hydrogen ions or protons (H+), moving down an electrochemical gradient, such as from the lumen into the
stroma of chloroplasts or from the inter-membrane space into the matrix in mitochondria.
The overall structure of ATP synthase is similar in all cells (Fig. 4a).
ATP synthase consists of two structural regions: the F0 domain within the membrane, which is involved in translocation of protons down their electrochemical gradient, and the F1 domain above the membrane and inside the matrix of the mitochondria, which contains catalytic sites responsible for ATP synthesis (Allegretti et al. 2015). In bacteria, the F0 and F1 regions are composed of ab2c10–14 and subunits, respectively. The c-subunits of the F0 domain form a ring that interacts with the a-subunit. The γ-subunit forms a central rotor stalk connecting to the c-ring at its base and penetrates
Figure 4. Rotational motors. (a) A 3D reconstruction of F-type ATP synthase dimer at 7.0 Å resolution by cryo-electron microscopy (EMDataBank entry: EMD-2852) (Allegretti et al. 2015).
The F-type ATP synthase was found to form a dimeric complex on mitochondrial membranes.
(b) 3D reconstruction of V-type ATP synthase at 9.7 Å by cryo-electron microscopy (EMDataBank entry: EMD-5335) (Lau and Rubinstein 2012). (c) 3D model of bacterial fl agellar rotor with its basal architecture highlighted by two orthogonal views (EMDataBank entry:
EMD-1887) (Thomas et al. 2006).
into the F1 catalytic domain. The F1 domain is assembled from three α- and three β-subunits arranged alternately in the form of a hexagonal cylinder around the γ-subunit. The peripheral stator stalk consists of 2 σ-subunits, with the σ-subun it binding to the F1 domain and 2-subunit anchoring the F0 domain in the mitochondrial membrane and interacting with the a-subunit. To use the energy of the transmembrane proton gradient to fuel ATP synthesis, the a- and c-subunits control proton transport in such a way that the c-ring rotates relative to the a-subunit, converting the energy of the electrochemical proton gradient into mechanical rotation of the c-subunits; the γ-subunit of the central stalk rotates with the c-ring and couples the transmembrane proton motive force over a distance of 10 nm to the F1 domain. The mechanical energy of rotation is used to release ATP, whose synthesis is catalyzed by the β-subunits in the F1 domain. Rotation of the c-ring and the central γ-subunit relative to the 33 subdomain is essential for coupling the proton motive force across the membrane to drive ATP formation and release. Since each c-subunit carries one proton, there are 10–14 protons transported per complete revolution of the c-ring, and roughly four protons are transported per ATP synthesized. The ATP synthase converts electrochemical energy to mechanical energy and back to chemical energy, with nearly 100% effi ciency (Wang and Oster 1998). ATP synthesis can occur at a maximal rate on the order of 100 sec–1 and sustains millimolar ATP concentrations in cells.
In some bacteria, the ATP synthase works in the reverse direction, in which the energy released by ATP hydrolysis drives the translocation of protons out of the cell, generating a proton gradient across the cytoplasmic membrane (Noji et al. 1997). Another family of protein motors that are closely related to the F0F1- ATP synthases is the H+-ATPases, which function in the direction opposite to ATP synthase in mitochondria, chloroplasts and bacteria (Fig. 4b) (Lau and Rubinstein 2012). H+-ATPases transport protons from the cytosol to the vesicle lumen or to the extracellular space through the membrane integral V0 domain. The cytoplasmic V1 domain couples the free energy of ATP hydrolysis to proton transport. The energy of ATP hydrolysis by the V1 domain is translated into rotation of the c-ring and proton transport. The a-subunit of the V0 domain transports protons via the nine transmembrane segments located in its C-terminal region. A positively charged arginine residue in the a-subunit stabilizes a negatively charged glutamate residue in one of the c-subunits prior to protonation.
The a-subunit uses two hemichannel structures to transport a proton and protonates the glutamate residue reversibly, which would release it from its interaction with the arginine residue in the c-subunit. The electrostatic attraction of the arginine residue with the unprotonated glutamate residue of the next c-subunit would allow rotation of the c-ring, bringing each c-subunit in contact with the a-subunit. This mechanism allows for ATP hydrolysis to drive unidirectional proton transport.
Another prominent rotatory motor is the bacterial fl agellar motor, powered by proton motive force generated by the proton fl ux across the bacterial cell membrane (Block and Berg 1984, Thomas et al. 2006). The rotation is driven by an ensemble of torque-generating units containing the proteins MotA and MotB (Fig. 4c). The rotor is consecutively rotated in the process of proton transport across the membrane. The rotor can rotate at a speed of 6000–17000 rpm alone, but may reach only 200–1000 rpm with the fl agellar fi lament attached. Like ATP synthase motor, the fl agellar rotor uses free energy with high effi ciency, generates power output of 1.5 × 105 pN nm s–1 and is the nanosclae engine driving the locomotion of bacteria in solutions (Ryu et al. 2000).
3.3 Locomotive Motors
There are two families of linear molecular motors, kinesins and dyneins, which move on microtubule rails, powered by ATP hydrolysis (Vale et al.
1996, Kon et al. 2012, Gee et al. 1997, Shingyoji et al. 1998). Kinesins usually move toward the plus ends of microtubules, and dyneins move toward the minus ends of microtubules. Both families of motor proteins share a common architecture: the motor head domain (Fig. 5a and 5b), which binds microtubules and generates force, and the tail domain, which binds cargo or membrane. The affi nity that the head binds to the microtubule is regulated through binding of ATP and ADP to the head domain. For kinesin, binding to a microtubule is the strongest when ATP is bound. By changing the strength of kinesin’s hold on a microtubule, ATP hydrolysis and nucleotide release regulate the attachment of the motor to the microtubules.
ATP hydrolysis also causes a conformational change in the head domain, which is amplifi ed to generate a larger movement of the whole motor molecule. Therefore, cycles of ATP hydrolysis and nucleotide release a couple of microtubule attachments with changes in the conformation of the motor’s head domain. Through this mechanism the motor steps along the microtubule in a head-over-head fashion, taking one step of 8 nm for each ATP hydrolyzed (Schnitzer and Block 1997). The two heads of kinesin work in tandem, as it walks along a microtubule. At the beginning of a walking cycle, one head is tightly bound to the microtubule and has no nucleotide in its active site. The second head has ADP in its active site and is positioned behind the fi rst head. Kinesin is then ready for its fi rst step and the coordination between the two heads takes place. ATP binding at the forward head (head 1) causes its neck linker to swing forward toward the plus end of the microtubule. Head 2 then moves to the leading position over the next binding site in the microtubule. It binds weakly and releases its ADP. ATP hydrolysis at head 1, then strengthens the interaction between head 2 and the microtubule, resulting in an intermediate with both heads