P2 receptor-mediated modulation of neurotransmitter release—an update

Presynaptic nerve terminals are equipped with a number of presynaptic auto- and heteroreceptors, including ionotropic P2X and metabotropic P2Y receptors. P2 receptors serve as modulation sites of transmitter release by ATP and other nucleotides released by neuronal activity and pathological signals. A wide variety of P2X and P2Y receptors expressed at pre- and postsynaptic sites as well as in glial cells are involved directly or indirectly in the modulation of neurotransmitter release. Nucleotides are released from synaptic and nonsynaptic sites throughout the nervous system and might reach concentrations high enough to activate these receptors. By providing a fine-tuning mechanism these receptors also offer attractive sites for pharmacotherapy in nervous system diseases. Here we review the rapidly emerging data on the modulation of transmitter release by facilitatory and inhibitory P2 receptors and the receptor subtypes involved in these interactions.
Introduction
Ionotropic P2X receptors and metabotropic P2Y receptors are the principal cell surface proteins, conveying the action of extracellular ATP, the ubiquitous signaling substance. P2X receptors are ligand-gated cation channels, composed of three individual subunits, whereas P2Y receptors belong to the superfamily of G protein-coupled receptors (GPCRs), with seven transmembrane domains. Various subtypes of P2X and P2Y receptor families are expressed throughout the brain and involved in a wide array of functions from fast synaptic transmission to long-term plasticity and trophic changes important for development, neuron-glia interactions, and neuroimmunomodulation. In addition, ATP modulates synaptic transmission pre- and postsynaptically, both in a positive and negative direction via activation of P2X and P2Y receptors, respectively.
The presynaptic nerve terminal is an important regulatory site, whereby the efficacy of synaptic transmission could be locally and efficiently controlled. Accordingly, axon terminals in the central nervous system and in the periphery are equipped with a wide variety of auto- and heteroreceptors [1–4]. Whereas presynaptic metabotropic receptors convey negative feedback regulation of transmitter release, presynaptic ionotropic receptors could amplify synaptic transmission. Moreover the activation of ligand-gated cation channels with high Ca2+ permeability could directly elicit transmitter release triggered by the Ca2+ influx through the receptor-ion channel complex [2, 4]. Coactivation of different presynaptic receptors provides a fine-tuning mechanism whereby different neurotransmitters and modulators can mutually influence the activity of each other. Presynaptic and extrasynaptic receptors controlling transmitter release also offer attractive target sites for existing and future pharmacotherapy, as they may modify the normal and pathological synaptic information processing without all-or-none actions [3].
Since ATP and its related nucleotides are ubiquitous signaling molecules, it is not surprising that their receptors, i.e., ionotropic P2X and metabotropic P2Y receptors, participate both in the negative and positive feedback modulation of neurotransmitter release. Although the principal function proposed for ATP-sensitive P2 receptors was that they mediate the fast transmitter action of extracellular ATP in neuro-neuronal and neuro-effector synapses in the nervous system, it was already recognized in the early 1990s that they are also involved in the regulation of transmitter release [5, 6]. It was subsequently revealed that the release of the major neurotransmitters of the brain and the peripheral neurons [acetylcholine (ACh), noradrenaline (NA), dopamine (DA), serotonin, glutamate, γ-aminobutyric acid (GABA)] are modulated by P2X and/or P2Y receptors. In 2000, Cunha and Ribeiro reviewed the literature on the presynaptic modulator role of ATP and suggested that there is a mismatch between the abundance of P2 receptor expression, the robust release of ATP in almost all parts of the central nervous system (CNS) and peripheral nervous system (PNS), and the relative paucity of identified P2 receptor-mediated synapses, which implicates the major role of ATP as a neuromodulator, rather than a classic transmitter [7].
The focus of this mini-review is the facilitatory and inhibitory modulation of neurotransmitter release by different subtypes of P2X and P2Y receptors, irrespective of their localization, i.e., whether they are pre-, post-, or extrasynaptic.
Therefore, in addition to a brief summary of the determining factors of ATP availability in synapses, the structure, pharmacology, signal transduction, and distribution of P2X and P2Y receptors in the nervous system, available information on the release-modulating P2 receptors, and the receptor subtypes involved in these interactions will be detailed and updated.
Determining factors of ATP availability in synapses
The participation of ATP and related nucleotides in the regulation of neurotransmitter release presumes their accumulation in the extracellular space upon ongoing neuronal activity. Extracellular purine availability in the nervous system is basically determined by the balance of release and removal by enzymatic degradation and uptake.
Sources and stimuli that trigger ATP release
Since ATP is ubiquitous, all metabolically active cells of the nervous system provide a potential pool for its release. Therefore, besides the nerve terminals themselves, the cellular source of released purines participating in the modulation of neurotransmitter release could be any cell type located in contact with nerve terminals, i.e., astrocytes, microglia, and endothelia. A wide variety of stimuli are known to release ATP to the extracellular space, which could lead to purine levels sufficiently high to activate nucleotide receptors expressed on the surface of pre- and postsynaptic membranes [8, 9]. Although the stimulation-dependent release of ATP upon conventional [10, 11] and high-frequency (e.g., [12]) neuronal activity is well documented, these stimuli probably result in a spatially restricted, localized increase in extracellular purine levels, which serve the fast synaptic transmission and its modulation within the synaptic cleft. Furthermore, ATP-metabolizing ectoenzymes, present on the nerve terminal membrane, and glial cells [13], such as ectoNTPDases, and the CD39/ecto-5′nucleotidase [14], may strongly limit nucleotide availability under these conditions. On the other hand, pathological events are known also to stimulate purine release. These signals include mechanical [15–17], chemical [18], and hypotonic stimuli [19], hypoxia/hypoglycemia/ischemia and consequent energy deprivation [20–25], inflammatory signals, such as bacterial lipopolysaccharide (LPS) [26, 27], interleukin-1β (IL-1β) [28], and cellular injury. The pathological ATP release might result in a purine-rich extracellular milieu leading to a more widespread activation of receptors reaching also the extrasynaptic receptors on the neighboring nerve terminals or distant cells such as astrocytes. Therefore, P2 receptors could play a role in the modulation of not only neuronal but also astrocytic transmitter release. Finally, nucleotides and nucleosides may promote further release of purines, by a homo- or heteroexchange mechanism, if they reach a relatively high concentration in the extracellular space [29].
Mechanisms of ATP release
Since ATP is a highly polarized molecule, which cannot pass freely the cell membrane being released to the synaptic cleft, it could enter the extracellular space by the following mechanisms: (1) vesicular exocytosis, (2) carrier-mediated release, (3) release through channels and membrane pores, and (4) cytolytic release. (1) Vesicular exocytosis is a prototype mechanism for neurotransmitters and neuromodulators to enter the extracellular space, which is expected to be a [Ca2+]o-dependent process. Indeed, ATP is taken up and stored in synaptic vesicles of nerve terminals [8] and astrocytes [15] and [Ca2+]o-dependent ATP release in response to neuronal stimulation appears in many areas of the central and peripheral nervous system (for further references see [8, 9, 24, 30, 31]). Moreover, recent findings indicate that vesicular ATP could be released not only from nerve terminals but also from neuronal somata [32] and astrocytes [15]. (2) Although specific transporters capable of transmembrane movement of ATP are yet to be molecularly identified in the nervous system, ABC (ATP binding cassette) proteins have been implicated as ATP transporters [19, 33, 34] in non-neuronal cells. These transporters are also expressed in glial cells [35] and mediate ATP release upon hypo-osmotic challenge [36, 37]. (3) Channels and pores, such as connexin hemichannels [38], are also potential candidates to drive the transmembrane movement of ATP. They have been identified to mediate ATP release from astrocytes and other non-neuronal cells in response to mechanical stress [39, 40] and other stimuli [31, 41]. (4) Although only scarcely supported by direct experimental proof [42], the general assumption is that any kind of cellular injury could result in high local ATP concentrations in the extracellular space. In this case the millimolar cytoplasmic ATP is expected to leak out of the cell through the membrane damage.
Metabolism of ATP in the extracellular space
Several enzyme families are responsible for the extracellular degradation of ATP in the nervous system. The first step of the inactivation of ATP is mediated by the family of ectonucleoside triphosphate diphosphohydrolases (ENTPDases, EC 3.6.1.5, also known as ectoATPase or apyrase), which are able to hydrolyze ATP and adenosine diphosphate (ADP) to AMP [14]. Among these enzymes ENTPDase 1, 2, 3, and 8 are present in the brain [43, 44], having low micromolar Km for ATP and ADP giving rise to rapid and highly effective hydrolysis of ATP in almost all neuronal tissues. In addition to the ENTPDase family, ATP and other nucleotides could also be dephosphorylated by ectonucleotide pyrophosphatases (ENPPs) and by alkaline phosphatases, both having broader substrate specificity, but also widespread tissue distribution [14]. The final step of extracellular inactivation is the hydrolysis of AMP by the ecto-5′-nucleotidase (EC 3.1.3.5) enzyme, which is the rate-limiting step giving rise to the formation of adenosine that acts on P1 receptors, which include A1, A2A, A2B, and A3 receptor subtypes. Thus, endogenous ATP is converted to adenosine to activate A1 adenosine receptors within a second in the hippocampus [45, 46], whereas the hydrolysis of ATP seems slower in other brain regions, such as the cerebral cortex [47]. Ectoenzymes therefore have an important role in the substrate delivery to different subtypes of P2X and P2Y receptors. In spite of its short half-life, effective concentrations of nucleotides can be reached in the synapse for the activation of ionotropic P2X receptors and metabotropic P2Y receptors.
Structure, pharmacology, and signal transduction of P2X receptors
Ionotropic P2X receptors are nonselective cation channels consisting of at least three subunits. P2X receptor subunits are 379-595 amino acid long polypeptide chains, having two transmembrane domains (TM1 and TM2) and a large extracellular loop [48, 49]. Until now seven members of this receptor family have been identified molecularly, which are numbered from P2X1 to P2X7, and have individual kinetics and pharmacological phenotype [50]. These receptor proteins coassemble into various homo- or heterooligomeric assemblies to form functional receptors. Among possible combinations so far 16 variations have been proved to be functional [51]. These are all of the homooligomeric receptors, except P2X6, which does not function in homooligomeric form, and the rest are heterooligomers, formed from P2X1-P2X6 subunits. However, recently it has been reported that by N-glycosylation even the homomeric P2X6 receptor could be rendered functional [52]. On the other hand, the P2X7 receptor functions only in homooligomeric form and does not coassemble with other known P2X receptor subunits. P2X receptors are permeable to both monovalent (Na+, K+) and divalent (Ca2+) cations and the activation of the receptor generates an inward current leading to the local depolarization of the cell membrane; in addition, the Ca2+ influx through the receptor-ion channel complex could directly trigger transmitter release. Moreover, upon prolonged or repetitive agonist application certain P2X receptors, especially the P2X7 receptor, display pore dilation which makes the channel permeable to high molecular weight cations up to 800 Da.
Basically P2X receptors are sensitive to ATP and to its various synthetic analogues but not to AMP and adenosine and the ligand binding profiles of homomeric P2X receptors are well established (for further information, see [50, 53]). On the other hand, less is known about the pharmacology of heteromeric receptors; among them, the pharmacological profile of P2X2/3, P2X2/6, P2X1/2, P2X1/4, P2X1/5, and P2X4/6 are described [54–59]. However, the expression pattern and the ligand binding profile of individual assemblies of P2X receptors are highly overlapping, often creating difficulties in the identification of the P2X receptor subunit composition of receptors expressed in native tissues. Moreover P2X receptor assemblies also share common ligand binding properties with certain members of the P2Y receptor family. Therefore, transgenic mice genetically deficient in individual P2X receptor subtypes are being increasingly used for P2X receptor identification.
The distribution of P2X receptors in neuronal structures
In situ hybridization studies with specific riboprobes, and immunocytochemical studies using antibodies raised against individual P2X receptor subunits, revealed that all seven P2X receptors are widely expressed in the nervous system. However, the expression of different receptor subunits show species-, region-, and cell type-specific distinct distribution [60]. Among the P2X receptors, P2X2, P2X4, and P2X6 seem to be most abundantly expressed in the brain, whereas other subunits show more restricted localization [60–63]. The typical localization of the P2X2 receptor is on nerve terminals of the brain and the periphery [61, 64, 65], although it also appears postsynaptically [62]. P2X1 receptors had initially been suggested to be exclusively expressed on smooth muscle membrane consistent with its role in mediating fast synaptic transmission at the autonomic neuroeffector junction [60]. However, recent studies with more sensitive probes revealed that its expression is more widespread, i.e., it is also present on central and peripheral neurons [63, 66]. The same holds true for P2X3 receptors, which are primarily associated with sensory pathways, but functional studies indicate that they are also expressed in other brain regions and autonomic pathways [67–69]. The P2X4 receptor shows substantial expression in several brain areas such as the cerebral cortex, hippocampus, thalamus, and brainstem [70] and is associated with postsynaptic specialization of synaptic contacts [62]. P2X5 subunits have the most restricted localization in the brain, although it shows strong representation in certain areas, e.g., the nucleus tractus solitarii (NTS) [71]. Finally, P2X7 receptors are also expressed in the brain, especially in reactive microglia and astroglia [72] and immunoelectron microscopic studies revealed a widespread presynaptic expression of P2X7 receptor immunoreactivity in a number of different brain areas, including the brainstem, hippocampus, cortex, spinal cord, and the skeletal neuromuscular junction [73–75]. However, two studies, using the same antibodies, demonstrated that P2X7 receptor immunoreactivity is still observable in the brain of P2X7 receptor knockout animals and thereby raised doubts on the validity of previous immunocytochemical observations [76, 77]. Therefore, available P2X7 receptor antibodies either recognize a site, which is not the P2X7 receptor, or, as a recent study indicates [78], a brain analogue of the P2X7 receptor, which shares its antibody binding domain with the cloned P2X7 receptor and partially retains its functionality in P2X7 receptor knockout animals.
Structure, pharmacology, and signal transduction of P2Y receptors
P2Y receptors all belong to G protein-coupled receptors, having seven hydrophobic transmembrane domains, and possess their ATP binding site on the external side of TM3 and TM7 domains [79–82]. The P2Y receptor family has eight individual members, numbered P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14. P2Y receptors are basically activated by adenine and uridine nucleotides, such as ATP, ADP, uridine diphosphate (UDP), and uridine triphosphate (UTP), but not by nucleosides and they are classified according to their sensitivity to purines and/or pyrimidines: P2Y1, 12 and P2Y13 are adenine nucleotide-preferring receptors; P2Y6 is preferred by uridine nucleotides; P2Y2, 4 and P2Y11 are receptors with mixed selectivity; whereas P2Y14 is activated by UDP-glucose, UDP-galactose, UDP-N-acetylglucosamine, and UDP-glucuronic acid [83]. Although a minority of P2Y receptor subtypes are incompletely characterized and the pharmacological profiles of individual P2Y receptors are partially overlapping, ligands are available which display some selectivity to certain subtypes of the P2Y receptor family [for further information see 84, 85]. Nevertheless, identification of individual P2Y receptors requires careful pharmacological analysis and the use of receptor knockout animals, if available, and exclusion of the involvement of adenosine receptors in the effect of nucleotides used as P2Y agonists, because they may be metabolized to adenosine (see, e.g., [45, 47]). In addition, the potential heteromerization of P2Y receptor subtypes with each other and with A1 adenosine receptors [86] should also be taken into account when individual P2Y receptors are identified.
Whereas P2X receptors convey rapid changes in the neuronal excitability on the millisecond timescale, P2Y receptors act on a longer, second timescale, appropriate for the fine-tuning of synaptic transmission. As for the signal transduction pathways activated by various subtypes of P2Y receptors, P2Y1, P2Y2, P2Y4, P2Y6, and P2Y11 receptors are coupled via Gq/11 proteins to stimulate phospholipase C, followed by increases in inositol phosphates and mobilization of Ca2+ from intracellular stores; in addition, the P2Y11 receptor mediates an increase in adenyl cyclase activity [84]. On the other hand, P2Y12, P2Y13, and P2Y14 receptors are coupled via Gi/o proteins to inhibit adenyl cyclase activity followed by a decrease in intracellular cAMP levels [84]. The activation of Gi protein-coupled P2Y receptors leads to the voltage-dependent inhibition of N-type voltage-sensitive Ca2+ channels directly or indirectly and subsequent inhibition of neurotransmitter release [85]. The inhibition of voltage-sensitive Ca2+ currents has also been demonstrated for those P2Y receptors, which are coupled to the Gq/11 proteins [85]; however, this inhibition is voltage independent [85]. In addition, the Gi/o-coupled P2Y receptors are known to activate voltage-sensitive GIRK K+ channels via direct interaction of Kir3 channel protein [85], which hyperpolarizes the neuronal membrane.
The distribution of P2Y receptors in neuronal structures
mRNA encoding all known P2Y receptors, i.e., P2Y1, P2Y2, P2Y4, P2Y6, P2Y12, P2Y13, and P2Y14 are present in the brain [81, 87–89]. Although our knowledge of their cell-specific localization at the protein level is still incomplete, it appears that a number of them, such as P2Y1, P2Y2, and P2Y6 receptors, are expressed both on neurons and astrocytes [90–96], whereas others are not exclusively, but predominantly localized to astrocytes (P2Y13: [97] P2Y14: [98]), oligodendrocytes (P2Y12: [99]], or microglia (P2Y12: [100]). P2Y receptor mRNA and protein can be detected in a number of different structures, including sympathetic and parasympathetic and sensory nerve terminals, basal ganglia, brainstem, cerebellum, cortex, hypothalamus, and hippocampus (for further references, see [85]). However, immunocytochemical data should be handled with caution due to the lack of verification of the specificity of many of the available antibodies. For detailed information on the distribution of individual P2Y receptor mRNAs and proteins, we refer to recent reviews on this particular topic [85, 101].
Modulation of neurotransmitter release by facilitatory P2 receptors
ACh
PNS
P2X receptors Since P2X receptors have relatively high Ca2+ permeability [102, 103], this property makes them capable of initiating neurotransmitter release by Ca2+ influx through the receptor-ion channel complex or facilitating Ca2+-dependent neurotransmitter release, provided that they are located nearby the release sites (Table 1). The first report suggesting that P2 receptors are involved in the facilitatory modulation of neurotransmitter release stems from 1991 when we found that opposite to the well-known inhibitory action of adenosine, α,β-methylene ATP, a metabolically stable analogue of ATP, enhanced electrically evoked acetylcholine release from the myenteric plexus of guinea pig and facilitated the related contractile response [6]. This effect was not blocked by the antagonists of adenosine receptors, and therefore was proposed to be mediated by ATP-sensitive P2 receptors [6]. In the same time Fu and Poo observed that ATP potentiates the spontaneous secretion of acetylcholine from developing neuromuscular synapses in Xenopus cell culture by promoting Ca2+ influx through the plasma membrane. However, this effect was not recognized to be the result of P2 receptor activation [104]. Later on, P2 receptor-mediated facilitation of acetylcholine release was confirmed by electrophysiological recordings in chicken ciliary ganglion [105] and mouse motor nerve terminals [106]. Presynaptic P2X receptors, involved in the facilitation of acetylcholine release, have also been identified in developing and adult neuromuscular synapses of Xenopus [107] and rat [108]. Homomeric P2X7 receptors are inserted into the membrane of mouse motor nerve terminals and their activation elicits vesicular exocytosis [74, 109]. However, there is no report about the presence of other subunit compositions of P2X receptors at the neuromuscular junction and it is also unclear whether such facilitatory receptors also exist on the terminals of central cholinergic neurons.
Table 1Facilitatory modulation of neurotransmitter release in the PNS and CNSPreparationMeasured effectProposed receptorReferenceAcetylcholine Guinea pig ileumNT quantificationP2[6] Chicken ciliary ganglionPresynaptic currentP2X[105] Xenopus NMJsEPP/mEPPP2[104, 107] Mouse NMJEPP/mEPPP2[106] Mouse NMJVesicular destaining, EPSCP2X7-like[74, 109] Rat NMJNT quantificationP2X[108]Noradrenaline Rabbit ear arteryEJPP2[110] PC12 cellsNT quantificationP2[111] Guinea pig ileumNT quantificationP2[6] Rat vas deferensNT quantificationP2X1, P2X3, P2X2/3[113] Guinea pig atriumNT quantificationP2X3, P2X2/3[114, 115, 118] Human and porcine heartNT quantificationP2X[119] Sympathetic neuronsNT quantificationP2X2[116] LC neuronsAP dischargeP2X[120] Rat hippocampusNT quantificationP2X1, P2X3[68]Serotonin Rat hippocampusNT quantificationP2[121]Dopamine Rat striatumNT quantificationP2Y[122, 123] Rat nucleus accumbensNT quantificationP2[124–128]Glutamate Rat brainstemmEPSC/NT quantificationP2X1[131, 157] Rat NTSEPSCP2X3, P2X2/3[129, 130, 132] Rat hippocampusNT quantificationP2X1, P2X3, P2X2/3[69] Rat hippocampusNT quantificationP2X7[75] Mouse hippocampusNT quantificationP2X7[137] Rat hippocampusEPSCP2X2[133] Rat hippocampusEPSCP2X7[134] Rat hippocampal neuronsEPSCP2[135] Rat cortical synaptosomesNT quantificationP2X7[136] Cultured astrocytesNT quantificationP2X7[142] Retinal Müller glial cellsNT uptakeP2X7[143] Rat spinal cordEPSC/mEPSCP2X3, P2X1/5, P2X4/6[144–148] Rat nucleus accumbensNT quantificationP2[149] Rat medial habenulaEPSCP2Y4[150] Cultured astrocytesNT quantificationP2Y1[151] Cultured Schwann cellsNT quantificationP2[152]GABA Midbrain synaptosomesNT quantificationP2X3, dinucleotide R[153] Cultured dorsal horn neuronsIPSCP2X[154] Cultured hippocampal cellsIPSCP2[156] Cultured cortical cellsNT quantificationP2X7[155] Rat brainstemIPSCP2X1[157] Rat, mouse, and guinea pig hippocampusNT quantification/IPSCP2X7 (indirect)[75, 137, 158] Cultured astrocytesNT quantificationP2X7[159] Rat hippocampusIPSCP2Y1[163, 206]Glycine Rat dorsal hornIPSCP2X[161] Rat trigeminal nucleussIPSCP2X[162]AP action potential, EJP excitatory junction potential, EPP end plate potential, EPSC excitatory postsynaptic current, IPSC inhibitory postsynaptic current, mEPP miniature EPP, mEPSC miniature EPSC, NMJ neuromuscular junction, NT neurotransmitter, sEPP spontaneous EPP, sIPSC spontaneous IPSC
Monoamines (NA, serotonin, DA)
PNS
P2X receptors The presynaptic facilitatory action of ATP on noradrenergic transmission was described for the first time by Miyahara and Suzuki in rabbit ear artery [110]. It was followed by the demonstration of the facilitatory effect of ATP and its metabolically stable analogue α,β-methylene ATP on [3H]noradrenaline efflux in PC12 cells [111] and in the guinea pig ileum [6]. However, the knowledge on P2 receptors at that time did not allow the identification of P2 receptor subtypes involved in these effects. The issue has been reinvestigated and it was found that sympathetic nerve terminals are equipped with ionotropic P2X receptors, activation of which directly elicits or facilitates noradrenaline release elicited by nerve stimulation [1, 112–115] via a direct Ca2+ influx through the receptor-ion channel complex. The pharmacological phenotype of these receptors varies between species, between transmission sites of the sympathetic nervous system, and even between the somata and nerve terminals of an individual neuron. Thus, in cultured sympathetic neurons of the rat an α,β-methylene ATP-insensitive P2X2-like receptor was identified [116], whereas in the guinea pig right atrium we found that α,β-methylene ATP stimulates noradrenaline outflow and the pharmacological profile of the underlying receptor was similar to that of P2X3 or P2X2/P2X3 receptors, consistent with the expression of their mRNA in the sympathetic ganglia [115]. In another study the facilitatory P2X receptors involved in the modulation of noradrenaline outflow in the rat vas deferens were identified as P2X1, P2X3, or P2X2/P2X3 receptors [113]. In contrast, cultured mouse sympathetic nerve terminals do not seem to express facilitatory nucleotide-sensitive receptors [117]. Importantly, these receptors seem to be endogenously activated by ATP released in response to ongoing neuronal activity [115, 116] and by myocardial ischemia in the guinea pig [118], porcine, and human heart [119] and could contribute to ischemia-induced arrhythmia and ischemic heart dysfunction.
CNS
P2X receptors In the central nervous system, locus coeruleus (LC) neurons of the rat are equipped with ATP-sensitive P2X-like receptors, which facilitate the discharge of spontaneous action potentials [120]. Facilitatory P2X receptors have also been described in the noradrenergic axon terminals innervating the hippocampus, and the homomeric P2X1 and P2X3 receptors were identified as the most likely subunits responsible for this action [68].
P2Y receptors P2 receptors enhance the release of serotonin from the hippocampus [121] and that of dopamine from the striatum [122, 123], and the latter effects are thought to be mediated by P2Y receptors. However, the pre- or postsynaptic localization of receptors responsible for these effects were not clarified in these studies. The P2 receptor agonist 2-methyl-thio ATP releases dopamine from the nucleus accumbens through direct and indirect mechanisms [124–126] in vivo. Interestingly, P2 receptor activation-evoked dopamine release seems to play a role in the modulation of feeding behavior as P2 receptor antagonists inhibit feeding-induced dopamine release and concomitant behavioral changes after food deprivation [127, 128].
Excitatory amino acids (glutamate, aspartate)
CNS
P2X receptors In addition to ACh and monoamines, the release of excitatory amino acid transmitters is also modulated by presynaptic P2X receptors in the CNS, as demonstrated partly by neurochemical and partly by electrophysiological methods.Activation of P2X receptors elicits glutamate release in the brainstem [129–132], hippocampus [69, 75, 133–135], and cortical synaptosomes [136]. As for the underlying receptor subunits involved in these effects, P2X1 [69], P2X2 [133], P2X3, and P2X2/3 receptors [69, 130] as well as P2X7 [75, 132, 134, 136, 137] were identified. The involvement of P2X2 receptors [133] and P2X7 receptors [137] has been confirmed by the use of transgenic mice deficient in P2X2 and P2X7 receptors, respectively. Moreover the activation of P2X7 receptors not only elicits glutamate release but also permits the activation of other ligand-gated ion channels on the nerve terminals, such as α7 nicotinic receptors, as demonstrated recently in rat cortical synaptosomes [136]. The activation of a P2X7-like receptor promotes Ca2+ influx in cortical synaptosomes [138] and in isolated midbrain synaptic terminals [139] and activates p38MAP kinase enzyme in the hippocampus [140]. This latter effect seems to participate in the effect of ATP to elicit glutamate release as it was sensitive to the inhibition by the specific p38MAP kinase inhibitor, SB203580 [140]. Nevertheless, the exact mechanism whereby the P2X7 receptor and subsequent activation of p38MAP kinase enzyme leads to increased glutamate release awaits further investigation. In addition, it has been reported that the P2X7 receptor agonist BzATP depresses synaptic transmission at the mossy fiber-CA3 synapse [141] in a p38MAPK-dependent way. However, more recently the participation of P2X7 receptors in this latter effect has been disproved [76, 77]. In addition to nerve terminals, P2X7 receptor activation also elicits glutamate release from cultured astrocytes [142] and inhibits the uptake of glutamate in Müller glial cells of the retina [143].The activation of P2X receptors facilitates excitatory transmission in the spinal cord, releasing glutamate from primary afferent fibers terminating in lamina II [144–148] and lamina V [144]; these actions are mediated by P2X3, P2X1/5, and P2X4/6 receptors, respectively. Finally, the ability of the P2 receptor ligand 2-methyl-thio ATP to release glutamate has also been demonstrated in vivo in the dopamine-depleted nucleus accumbens [149], although the underlying receptor subtype was not identified in this study.
P2Y receptors Interestingly, the activation of P2Y receptors is also implicated to elicit and potentiate glutamate release in the central nervous system. In the medial habenula nucleus UDP and UTP increase presynaptic release probability and elicit a non-Hebbian-type long-term potentiation of excitatory transmission, an effect probably mediated by P2Y4 receptors [150]. In addition, the activation of P2Y1 receptors elicits vesicular glutamate release from astrocytes [151] and from cultured Schwann cells [152].
Inhibitory amino acids (GABA, glycine)
CNS
P2X receptors ATP or P1,P5-di(adenosine-5′) pentaphosphate (Ap5A) elicits an increase in the intrasynaptosomal calcium and induces subsequent GABA release in midbrain GABAergic synaptosomes via activation of P2X3 and a dinucleotide receptor [153]. The regulation of GABA release by P2X receptors has also been reported in the spinal cord [154], cultured cortical [155] and hippocampal [156] cells, and the brainstem, where the excitatory and inhibitory synaptic transmission is facilitated via P2X3 and P2X1 receptors, respectively [157]. In addition to direct modulation of glutamate release, P2X7 receptor activation also releases GABA from the hippocampus through the activation on non-NMDA-type glutamate receptors [75]. This effect is absent in mice genetically deficient in P2X7 receptors [137] and mediated by the sodium-dependent reversal of the GABA transporter [75]. P2X receptor-mediated, TTX-sensitive GABA release has been implicated in the accelerated recovery of guinea pig hippocampal slices from a hypoxic/hypoglycemic insult [158]. The activation of P2X7 receptors also releases GABA from cultured RBA astrocytes, however, with a different mechanism, by participation of the HCO3-/Cl- exchanger [159]. On the other hand, no evidence was found for a direct facilitation of GABA release by P2 receptors in the hippocampal nerve terminal preparation [160]. Nevertheless, the release of another inhibitory transmitter, glycine, is augmented by P2X receptor activation in the dorsal horn [161] and in the brainstem trigeminal nucleus [162].
P2Y receptors In addition to P2X receptors, activation of P2Y1 receptors leads to an increase of the inhibitory postsynaptic current (IPSC) frequency in an acute hippocampal slice in a manner dependent on action potential generation, indicating that this effect is related to the activation of receptors present on the somata/dendrites of hippocampal interneurons [163].
Modulation of neurotransmitter release by inhibitory P2 receptors
In addition to facilitatory modulation, P2 receptors are also involved in the inhibitory modulation of the release of various transmitters and the metabotropic P2Y receptors are thought to play a major role in these actions (Table 2).
Table 2Inhibitory modulation of neurotransmitter release in the PNS and CNSPreparationMeasured effectProposed receptorReferenceAcetylcholine Frog ganglionEPSCP2[165] Guinea pig ileumNT quantificationP2[164, 166, 167] Ileal synaptosomesNT quantificationP1[168] Guinea pig submucosal neuronsEJPP3[169] Frog NMJEPPP2[170] Rat submandibular ganglionEPSCP2[171] Rabbit retinaNT quantificationP2[172] Mouse NMJsEPPP2Y[173]Noradrenaline Mouse vas deferensNT quantificationP2Y-like[5, 174, 181, 182] Rat vas deferensNT quantificationP3[178] Rat vas deferensNT quantificationP2Y[181] Rat vas deferensNT quantificationP2Y12, P2Y13[113] Guinea pig vas deferensNT quantificationP2Y-like[179] Guinea pig saphenous arteryEJPP2[175] Rat caudal arteryNT quantificationP3, P2Y[176, 177, 180] Rat atriumNT quantificationP2Y-like183 Rat irisNT quantificationP2Y-like[184] Rat kidneyNT quantificationP2[185] Rat pancreasNT quantificationP2[186] Chick sympathetic neuronsNT quantificationP2, P2Y12[187, 188, 189] Bovine chromaffin cellsNT quantificationP2Y12[190, 194] Rat cortexNT quantificationP2Y[195] Rat hippocampusNT quantificationP2Y[196]Serotonin Rat cortexNT quantificationP2[197]Dopamine Rat striatumNT quantificationP2[198]Glutamate Rat cortexEPSCP2[199] Rat hippocampusEPSCP2Y[200, but see 201] Rat spinal cordPolysynaptic EPSPP2Y[204] Hippocampal slice cultureEPSCP2Y[205]EJP excitatory junction potential, EPP end plate potential, EPSC excitatory postsynaptic current, EPSP excitatory postsynaptic potential, NMJ neuromuscular junction, NT neurotransmitter, SEPP spontaneous EPP
ACh
PNS
It has been known for a long time that ATP is involved in the inhibitory presynaptic modulation of cholinergic transmission [164]. However, it has been the subject of a long-standing debate whether ATP itself is responsible for this effect or its degradation product adenosine [165–167], whereas an alternative was that ATP itself acts on adenosine receptors [168] or activates a putative P3 receptor bearing pharmacological features of both P1 and P2 receptors [169]. A more definitive proof for the involvement of P2 receptors in the inhibition of acetylcholine release was obtained later in the frog neuromuscular junction [170], rat submandibular ganglia [171], and rabbit retina [172]. Presynaptic P2Y receptors responsible for the inhibition of spontaneous acetylcholine release were recently identified at the mouse neuromuscular junction [173]. In this study the underlying subcellular mechanism of the inhibition of acetylcholine release was also explored: the activation of P2Y receptors is coupled to Gi/o proteins and modulates presynaptic Ca2+ channels related to tonic secretion of acetylcholine [173].
CNS
Inhibitory P2 receptors involved in the modulation of ACh release have been demonstrated in rat cerebral cortex [47]. By contrast, in the hippocampus, ATP primarily inhibits acetylcholine release through its breakdown to adenosine and subsequent action on A1 adenosine receptors [47].
Monoamines (NA, serotonin, DA)
PNS
The presence of nucleotide-sensitive inhibitory P2 receptors on postganglionic sympathetic neurons was recognized relatively early [5, 174–179], although initially these receptors were qualified as P2Y-like [5] or as putative P3 receptors, which are “hybrid” receptors between P1 and P2 receptors and sensitive to adenine nucleotides but also to theophylline derivatives [176–178]. These receptors have been described and characterized in sympathetic nerves innervating the rat caudal artery [176, 177, 180], guinea pig saphenous artery [175], vas deferens [5, 178, 179, 181, 182], atrium [183], iris [184], kidney [185], and pancreas [186] as well as in cultured sympathetic neurons [187, 188]. As for subtype-specific identification, Queiroz et al. [113] identified presynaptic inhibitory nucleotide receptors on the noradrenergic axon terminals of the rat vas deferens as P2Y12 and/or P2Y13 receptors, whereas on cultured sympathetic neurons [189] and bovine adrenal chromaffin cells [190] only P2Y12 receptors have been identified. Interestingly, it appears that mouse sympathetic neurons [191] and noradrenergic nerves innervating the rat adrenal cortex [192] do not express an inhibitory P2 receptor. The mechanism of P2Y receptor-mediated inhibition of noradrenaline release has also been explored in several studies: the activation of P2Y receptors inhibits voltage-dependent Ca2+ influx and thereby limits the Ca2+-dependent vesicular exocytosis and subsequent efflux of noradrenaline to the extracellular space [189, 193, 194].
CNS
Similar inhibitory P2Y receptors have also been reported in the CNS in the rat brain cortex [195] and hippocampus [196]; however P2Y receptor subtypes involved were not identified in these early studies.
In the CNS, ATP inhibits the release of serotonin [197] and dopamine [198] via activation of metabotropic P2 receptors.
Excitatory amino acids (glutamate, aspartate)
CNS
ATP and its metabolically stable analogue ATP-γ-S inhibits depolarization-evoked glutamate release from rat brain cortex slices [199] and inhibits glutamatergic EPSPs in hippocampal CA1 synapses [200]. Although the underlying receptor was sensitive to theophylline derivatives, the authors proposed that ATP acted through a putative pertussis toxin-sensitive P2Y receptor. However, this hypothesis was challenged by showing the rapid and highly effective hydrolysis of ATP in the hippocampal slices [45, 46] and by the demonstration of the complete absence of nucleotide-mediated modulation of excitatory synaptic transmission in the hippocampi of A1 receptor-/- mice [201]. In a recent study Rodrigues et al. demonstrated that single hippocampal pyramidal neurons do express P2Y1, P2Y2, and P2Y4 receptors, and the release of glutamate, measured by a neurochemical technique, is inhibited by these receptors [69]. The discrepancy between the observations obtained in electrophysiological and neurochemical studies might be explained by the fact that in the former, individual synapses, whereas in the latter, glutamate release from all synapses of the hippocampal slice were simultaneously investigated. Nevertheless, the exact conditions under which the activation of P2Y receptors by endogenous ligands gain significance remain to be identified.
Functional data suggest that the release of glutamate in the spinal cord is modulated by inhibitory P2Y receptors. The activation of P2Y receptors causes blockade of the N-type calcium channels in dorsal root ganglion (DRG) cells [202], and this effect may decrease the release of glutamate from DRG terminals in the spinal cord and thereby partly counterbalance the algogenic effect of ATP [203, 204]. This assumption is supported by the findings that the P2Y1/12/13 receptor agonist ADP-β-S inhibits polysynaptic, but not monosynaptic excitatory postsynaptic potentials in the hemisected spinal cord and exhibits antinociceptive potential in the tail flick test [204].
Recent studies revealed that modulators released from glial cells also regulate neurotransmitter release from nearby nerve terminals by the activation of P2 receptors. Hence mechanical stimulation of astrocytes in hippocampal cell culture leads to the generation of Ca2+ waves in astrocytes, which spread by the release of ATP and subsequent activation of P2 receptors and lead to the depression of excitatory synaptic transmission between neurons [205]. This glia-driven synaptic depression is partly mediated by ATP itself acting on P2Y receptors and partly by adenosine acting on A1 adenosine receptors [205]. A similar mechanism has also been demonstrated in intact hippocampal slices, where ATP released from neurons and astrocytes acts on P2Y1 receptors to excite interneurons, resulting in increased synaptic inhibition within intact hippocampal circuits [206]. On the other hand, to our knowledge there is no information regarding whether the release of GABA and other inhibitory amino acids is subject to modulation via inhibitory P2 receptors.
Potential therapeutic utilization of P2 receptors involved in the regulation of neurotransmitter release
P2X and P2Y receptors involved in the regulation of neurotransmitter release offer attractive, although not yet utilized sites for pharmacotherapy in nervous system diseases. For instance, facilitatory P2X receptors present on axon terminals could be activated not only during normal neuronal activity, but also during pathological situations, when cellular damage provides an ATP-rich extracellular milieu nearby the receptors. Thus, P2X receptors present on the sympathetic nerve terminals supplying the heart seem to be endogenously activated by ATP by myocardial ischemia [118, 119] and could contribute to ischemia-induced arrhythmia and ischemic heart dysfunction. Therefore, inhibition of these facilitatory P2X receptors might have therapeutic relevance in ischemic heart disease. The pathological activation of CNS P2X receptors, regulating the release of glutamate during ischemic-like conditions, was also recently described [207]. Increased activation of P2X receptors could contribute to ischemia-evoked glutamate release and thereby to glutamatergic excitotoxicity and resultant neuronal death; therefore, inhibition of these receptors could be a promising approach to treat ischemia-related neurodegenerative diseases. An analogous mechanism could play a role in the spinal cord during the sensitization process leading to various forms of sensory neuropathy; therefore, attenuation of increased glutamate release from the central terminals of primary sensory neurons by the inhibition of P2X receptors is a potential pathway which could be utilized in neuropathic pain. Since inhibitory P2Y receptors are frequently coexpressed on nerve terminals that are equipped with P2X receptors, activation of P2Y receptors could have a similar effect as the inhibition of P2X receptors. Therefore P2Y receptor agonists might also have therapeutic value in the areas described above. However, one should bear in mind that different subtypes of P2X and P2Y receptors affect various other aspects of physiological and pathological neuronal functions, which could also modify their potential.
Conclusion
In conclusion, substantial advances have been obtained in the identification and characterization of neurotransmitter release modulating P2 receptors in recent years. It appears that almost all major neurotransmitters of the nervous system are subject to neuromodulation by nucleotide-sensitive P2 receptors. Although there are exceptions to this rule, in general the release of different transmitters is subject to a dual modulation similar to modulation of other transmitters of the CNS and PNS: facilitatory modulation is conveyed by ionotropic P2X receptors, whereas inhibitory modulation is mediated by G protein-coupled metabotropic P2Y receptors. Amongst P2X receptors, P2X1, P2X2, P2X3, P2X2/3, P2X1/5, P2X4/6, and P2X7 receptors were identified to be responsible for facilitatory modulation in different areas of the CNS and PNS. In addition, P2Y receptors (P2Y1, P2Y4) could also mediate facilitation of transmitter release in certain areas. Inhibitory modulation of neurotransmitter release is mediated by P2Y12 and P2Y13 receptors; however, individual P2Y receptor subtypes involved in these interactions are far from fully explored yet. It appears that not only neuronal, but also glia-derived ATP play a role in the modulation of neurotransmitter release. The intensity of P2 receptor-mediated modulation, the balance between the facilitatory and inhibitory modulation and the participating individual receptor subtypes, however, varies between individual transmission sites, depending on the expression pattern of P2 receptors and the factors determining the nucleotide levels in the vicinity of release of modulatory P2 receptors. Therefore, further progress is necessary in order to obtain a precise mapping of P2 receptor-mediated modulation of neurotransmitter release. The in vivo relevance of most of the in vitro observations on presynaptic P2 receptors awaits further investigation. Finally, physiological and pathological situations where presynaptic P2 receptors become endogenously activated by released nucleotides need to be identified.

transmitter
release
presynaptic
p2y receptors
p2x receptor
neuromodulation


Optical molecular imaging for systems biology: from molecule to organism

The development of highly efficient analytical methods capable of probing biological systems at system level is an important task that is required in order to meet the requirements of the emerging field of systems biology. Optical molecular imaging (OMI) is a very powerful tool for studying the temporal and spatial dynamics of specific biomolecules and their interactions in real time in vivo. In this article, recent advances in OMI are reviewed extensively, such as the development of molecular probes that make imaging brighter, more stable and more informative (e.g., FPs and semiconductor nanocrystals, also referred to as quantum dots), the development of imaging approaches that provide higher resolution and greater tissue penetration, and applications for measuring biological events from molecule to organism level, including gene expression, protein and subcellular compartment localization, protein activation and interaction, and low-mass molecule dynamics. These advances are of great significance in the field of biological science and could also be applied to disease diagnosis and pharmaceutical screening. Further developments in OMI for systems biology are also proposed.
Introduction
Due to the rapid progress in the field of molecular biology that has occurred over the last century, we have come to see that life involves thousands of genes, proteins, metabolites, etc. However, life is not invoked by simply collecting these together; an integrated system formed from those components is required. It is vital to understand biological systems such as molecular networks, cells, tissues and even the organism itself at the system level. This shift in biological paradigm from reductionism to integration has made the field of systems biology [1–5] a new challenge that is currently a hot topic. Although attempts to systematically understand biological systems can be dated back to Wiener’s time [6], research into systems biology represent the first attempts to achieve this based on detailed knowledge of molecules. It promises to reveal the relationships among elements of systems that may include just a few proteins that together perform a defined task or more complex molecular machines, cells and groups of cells, with the goal of understanding their emergent properties [3].
It is widely agreed that systems biology is a large scientific field that relies greatly on collaborations between disciplines such as the life sciences, information science, system engineering and analytical technology. One of the keys to successful research in this field is considered to be work being done in analytical chemistry [7, 8] to develop innovative analytical methods that meet the needs of systems biology research. For example, capillary electrophoresis has been recognized as being the “gold standard” in genomics for DNA sequencing. Similar “gold standard” methods are available in transcriptomics (DNA microarray), proteomics (multidimensional separation coupling with mass spectrometry) and metabolomics (microseparation coupled with mass spectrometry and nuclear magnetic resonance). However, more of these methods are needed. As well as revealing genes, proteins and metabolites from those omic investigations, which relate to the basic structures of systems, it is important to have methods capable of monitoring their localizations, connections, and in particular their dynamics over time under various physiological or pathological conditions.
Optical molecular imaging (OMI) is a versatile technique that can be used to investigate the dynamics of biological events in molecules, cells, tissues and organisms in real time and in vivo (Fig. 1) [9–11]. Compared with other imaging approaches like magnetic resonance imaging and positron emission tomography etc., OMI exhibits the great advantages of high temporal (picosecond) and spatial (submicron) resolutions, high sensitivity (single-molecule level) and minimal invasion, and shows high potential for systems biology. This review highlights recent advances in OMI, focusing especially on the development of fluorescent probes such as fluorescent proteins and semiconductor nanocrystals (also referred to as quantum dots, QDs), OMI instrumentation and approaches (techniques). The application of OMI to studies of localization, conformations and interactions of biomolecules in vivo are also highlighted. In particular, this review describes the application of OMI for tracing the dynamics of single molecules and whole-body living organisms, which strongly suggest that OMI is an informative systems biology method that may be used to uncover biological events at different system levels. The future development of OMI is also predicted.
Fig. 1Utilization of OMI in life science research. OMI can investigate the dynamics of biological events in real time from molecules (left), cells, tissues and organisms (right) digitally and quantitatively. It can handle a wide range of intensities (about 12 orders of magnitude), times (femtoseconds to years) and spatial dimensions (nanometers to centimeters), and it gives high spatial and temporal resolution of the targeted cellular structures—better than any other method
Advances in fluorescent probes
Aequorea fluorescent protein (AFP)-derived mutants
Labeling of biomolecules with fluorescent probes or other dyes has facilitated in vitro or in vivo studies of biomolecular structures and dynamics, as well as their interactions, which is critical if we are to understand the biomolecular mechanisms of cellular function. However, traditional methods of chemical labeling based on fluorescent dyes are often inadequate for biomolecular labeling, repurification and reintroduction into cells by invasive methods like microinjection. These limitations have spawned efforts to noninvasively and site-specifically label protein in living cells or tissues by using green fluorescent protein (GFP) and its variants. The fluorescent protein presents a relatively small size and a compact, single-domain structure, which allow it to fuse to other targeted proteins with little or no interference in native protein.
GFP was cloned from Aequorea victoria. Excitation of the wild-type GFP by light with a wavelength of 398 nm can induce the emission of bright green fluorescence, peaking at 508 nm. Through continuous efforts to perform protein mutation of GFP, scientists have developed various kinds of AFP with different excitation and emission wavelengths, enhanced brightness, and improved pH resistance compared to the original wide-type, such as enhanced GFP (EGFP), cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP) (Fig. 2a) [12]. The original YFP exhibited several drawbacks when imaging in vivo [13]. Later generations of YFP mutants, including “Citrine” [14] and “Venus” [15], exhibit low pH (5.7) resistance, halide insensitivity, greatly improved photostability, brighter fluorescence, and faster maturation. The original generation of CFP mutants also displayed several spectroscopic disadvantages. By using similar methods to YFP mutagenesis, a CFP mutant named “Cerulean” [16] was obtained, with an improved quantum yield, a higher extinction coefficient and a longer fluorescence lifetime. Another new color GFP mutant, cyan-green fluorescent protein (CGFP) [17], whose excitation and emission wavelengths are intermediate between those of CFP and EGFP, was also found by using this tactic. In addition, “PA-GFP,” a GFP mutant, has 100 times greater fluorescence than the original and remains stable for days [18]. These fluorescent proteins were termed molecular or optical highlighters, and represent perhaps the most promising tools for investigating protein lifetimes, transport, turnover rates and so on.
Fig. 2a–cVarious FP mutants. a Introduction of the mutation of Thr203His in GFP results in significantly red-shifted maximum excitation and emission wavelengths; this mutant is named YFP. b By using mutagenesis, the original tetrameric DsRed is reconstructed into the monomeric DsRed variant. c Interchanging the amino and carboxyl portions of GFP and rejoining them with a short spacer generates cpGFP
Other kinds of fluorescent protein
Long-wavelength fluorescence can provide greater tissue penetration and better spectral separation from cellular autofluorescence. Therefore, red fluorescent proteins (RFP) are very useful in applications related to multicolor protein-tracking and the construction of sensors. The first RFP with a red emission wavelength was cloned from Discosoma genus, named “DsRed” [19]. Its structure is similar to GFP, but it is tetrameric. By using mutagenesis, different derivations exhibiting several advantages have been generated, such as the fluorescent timer of DsRed which has fluorescence that changes from green to red over time [20], the monomeric DsRed, named “mRFP1” [21] (Fig. 2b), as well as DsRed2 [22], T1 [23], and mCherry [24], which have chromophores that are brighter, more soluble, and mature faster than the original generation. Another orange-emitting fluorescent protein, which has higher pH-resistance and is more effective for imaging, was discovered in Fungia concinn [25]. In addition, “Kaede” [26], which is cloned from Trachyphyllia geoffroyi, can convert from green to red fluorescence and brightens 2,000-fold under UV illumination.
New rearrangement variants of fluorescent protein
In previous studies, fluorescent proteins were treated as an indivisible entity, usually appended to the amino or carboxyl terminus of the targeted protein. However, several rearrangements of GFPs in which the amino and carboxyl portions were interchanged and rejoined with a short spacer still remained fluorescent (Fig. 2c). These circular permutations exhibit several advantages, such as altered pKa values, orientations of the chromophore with respect to a fusion partner, and greater absorbance of the exciting energy [27, 28].
Nongenetic fluorescent probes
However, there are some disadvantages of using FPs to perform OMI of the whole-body. Given the available wavelengths of excitation and emission and the power of the FP, optical imaging with a depth of penetration of approximately 1–2 mm limits their use to C. elegans, Drosophila or surface structures in small animals. To solve these problems, applying QDs (quantum dots) to OMI of the whole-body could allow us to visualize biological events in deeper tissues with better image quality.
Compared with conventional organic dyes and fluorescent proteins, semiconductor nanocrystals—quantum dots—exhibit a narrow, tunable, symmetric emission spectrum and better photochemical stability. Also, their wavelengths of maximum excitation and emission shift to shorter wavelengths with decreasing size [29, 30]. QDs present several great advantages for whole-body OMI, such as their very large molar extinction coefficients and their very bright emission, which mean that the emission can be visualized in big animals [31]; the ability to perform simultaneous multiple color imaging of targets of interest [32]; and long excited state lifetimes that provide a way to separate the QD fluorescence from background fluorescence [33]. Because QDs are capped with a monolayer of organic ligands and are hydrophobic, they cannot be used for visualization in vivo. Goldman et al. [34] reported that QDs capped with fusion proteins of a specific antibody, protein G and leucine zipper adaptor protein could be used to image the target protein in vivo (Fig. 3). Since then, bioconjugated QDs have provided new approaches for the ultrasensitive and multiple-color imaging of targeted molecules in vivo. In addition, integrating QDs with paramagnetic substances results in new multimodality imaging probes that cuple the deep imaging capability of magnetic resonance imaging with ultrasensitive OMI [35]; these would be very powerful probes if used in clinical research. More recently, a new QD conjugate was found to emit long waves in the absence of external excitation light through bioluminous resonance energy transfer [36]. Compared with existing quantum dots, self-luminous quantum dots provide great sensitivity when imaging small animals, giving high signal-to-noise ratios.
Fig. 3Method of conjugating QDs to target proteins. The pG-zb acts as a molecular adaptor, connecting the QDs with the target protein through interactions of its protein G portion with a specific antibody as well as interactions of its positively charged tail with QDs capped with a negatively charged dihydrolipoic acid surface
OMI techniques
General imaging microscopy
Laser scanning confocal microscopy (LSCM) (Fig. 4a) and wide-field microscopy (WFM) are the tools most widely used for in vivo OMI. The major difference between them is that LSCM only collects the fluorescence emission of in-focus light, whereas WFM collects all signals, including out-of-focus light [37]. LSCM can produce superior images of multiple-cell samples, with photobleaching. However, LSCM cannot perform multiple-color imaging because it only observes in-focus light [38]. WFM is among the most sensitive of all such methods, permits minimal exposure of the sample, uniform illumination, an unlimited choice of excitation wavelengths, and is relatively simple [37]. One of the disadvantages of WFM is that disturbances are seen in observations of thicker specimens, where the out-of-focus signals become substantial [38].
Fig. 4a–fFluorescent OMI approaches. a LSCM only collects in-focus emitted light. b The principle of multiple photon excitation is based on the use of pulsed long excitation wavelengths to excite fluorescence. c FRET occurs between a donor and an acceptor that are in molecular proximity if the emission spectrum of the donor overlaps the excitation spectrum of the acceptor. d FRAP can reveal the mobility of FP-labeling proteins. These images illustrate the change in fluorescence of cells expressed with YFP-hGR before and after photobleaching. Reproduced from [51] with permission. e FLIM can measure the time-dependent emission intensity. The histogram represents the fluorescence lifetime distributions for the donor in the presence of interactions (red) or not (yellow). Reproduced from [57] with permission. f FCS can monitor the fluorescence signals emitted from the ROI. The cross-correlation curve (black) indicates a higher level of dimer or oligomer formation in the R1- and R5-expressing cells. Reproduced from [59] with permission
Furthermore, multiple-photon laser scanning microscopy (MPLSM) has also been used in OMI [39]. The principle of MPLSM is that pulsed long excitation wavelength light, such as that provided by an infrared (IR) laser beam, is used to excite a molecule at the focal plane by multiple photons, causing fluorescence (Fig. 4b). The advantages of MPLSM are high tissue penetration ability (>30 μm), low photobleaching or photodamage, and a high signal/noise ratio (due to low autofluorescence) [40].
Second-harmonic imaging microscopy (SHIM)
SHIM is based on a nonlinear optical effect called second harmonic generation (SHG), commonly called frequency doubling. This phenomenon requires that intense laser light passes through a highly polarizable material with noncentrosymmetric molecular organization [41]. Biological macromolecules often assemble into large, ordered noncentrosymmetric structures and have highly polarizability. Therefore, high-resolution SHIM imaging can visualize living cells or tissues. This method provides several benefits: the targeted molecules are not excited, photobleaching does not occur, and it can detect information related to pathology. In addition, a combination of SHIM and MPLSM could make a powerful optical imaging technique.
Optical coherence tomography (OCT)
The principle of OCT imaging is analogous to that of ultrasound B-mode imaging, except that OCT uses light rather than acoustic waves [42]. OCT can provide cross-sectional imaging of structures below the tissue surface, analogous to histopathology. By using a state-of-the-art laser as the light source, ultrahigh-resolution imaging with axial resolutions as fine as 1–2 μm can be achieved [43]. However, the penetration depth of the technique in most tissues is limited to approximately 2–3 mm [44]. Several features of OCT imaging make it well-suited to imaging-based diagnostics and surgical guidance, such as its high resolution, its ability to quantitatively assess, and its ability to image some tissue functions.
Fluorescent molecular imaging approaches
Fluorescence resonance energy transfer (FRET)
FRET is a quantum mechanical phenomenon that occurs between a fluorescence donor and a fluorescence acceptor with a favorable dipole–dipole orientation that are in molecular proximity to each other, provided the emission spectrum of the donor overlaps the excitation spectrum of the acceptor (Fig. 4c) [45]. Therefore, imaging based on FRET can determine the proximity (within the nanometer range) between labeled biomolecules in living cells [46, 47]. FRET-based microscopy includes intensity-based detection methods and fluorescence decay kinetics-based detection methods [46]. Similar to general fluorescence imaging microscopy, FRET also suffers from various drawbacks, like autofluorescence, detector noise, optical noise and photobleaching. In addition, spectral bleed-through (SBT), or cross-talk, is a major problem in FRET [48]. FRET is currently widely used in studies of protein colocalization, conformational changes, protein interactions and signal transduction [47, 49].
Fluorescence recovery after photobleaching (FRAP)
FRAP involves observing the rate of recovery of fluorescence resulting from the movement of a fluorescent marker into an area which contains the same marker that has been rendered nonfluorescent via an intense photobleaching pulse of laser light (Fig. 4d). FRAP has proved to be a powerful method for measuring the mobilities of target molecules in various membranes, cytoplasms and nuclei [50]. For example, the mobility of the hormone receptor in the nucleus of a living cell [51], the mobility in subcellular compartments and the dynamics of vesicles inside the synapses of cultured hippocampal neurons [52] have been studied using this approach.
Fluorescence lifetime imaging microscopy (FLIM)
FLIM is a technique by which the mean fluorescence lifetime of a chromophore is measured at each spatially resolvable element of a microscope image [53]. The fluorescence lifetime is an inherent property of a chromophore that is sensitive to environmental and physical processes. Therefore, FLIM can detect interesting physical processes that can influence the excited state of a probe in vivo [54]. In addition, the mean lifetime of a donor is shifted to shorter lifetimes in the presence of an acceptor in FRET (Fig. 4e), so FLIM can detect changes in FRET [55], although this may not be valid in complex cellular systems that consist of several different lifetime components [56]. For example, using a FRET–FLIM method, the regulation of the activity of receptor protein tyrosine kinase (RPTK) through its dimerization was demonstrated in living cells [57]. The main drawbacks of FLIM are that it has relatively low sensitivity and requires expensive instrumentation.
Fluorescence correlation spectroscopy (FCS)
FCS is an approach that has emerged only recently. It is a high-sensitivity photon-counting technique that permits fluctuations in the fluorescence signal due to changes in the fluorescence quantum yield to be measured (Fig. 4f) [58]. Therefore, FCS can directly measure the mobilities of biomolecules, and it can monitor the average association and dissociation of target labeled with fluorescence probe—which vary with interaction kinetics, complex composition and complex size—in vivo without photobleaching. For example, FCS can be used to detect the clustering of somatostatin receptors [59] and the binding affinities of glucocorticoids (GC) in different subcellular compartments [60].
Applications of OMI
OMI for biomolecules
Originally, OMI of biomolecule function was performed on dead fixed cells or tissue sections in vitro. The first application of fluorescent detection in situ emerged in 1980, when RNA that was directly labeled with fluorophore was used as a probe for specific DNA sequences [61]. However, this method is not sensitive enough. Another indirect detection method allows signal to be significantly increased by binding secondary reporter to the hybridization probes that bind to targeted DNA [62] and mRNA [63]. Recently, Santangelo et al. [64] indicated that coinjection of two beacons based on FRET could be used to map the localization of a specific hybridization in living cells. Another application of FISH is immunocytochemistry (IC), which is used to detect target protein in specimens in vitro as antigen by means of antibodies labeled with fluorescence probe [65]. The technique of FISH is still being developed, with more specific fine-tuning of sensitivity and multiplicity necessary, after which its applications should gradually move from being in vitro to in vivo [66].
In addition, Fourier transform infrared (FT–IR) spectroscopy allows the simultaneous detection of nearly all organic molecules in a single spectrum and hence is ideally suited for the investigation of complex metabolic pathways. However, this advantage is only useful when powerful mathematical methods are available for the analysis of the highly complex spectra produced. Glycolytic intermediates In yeast extract were identified with FT–IR spectroscopy. The results were in good agreement with known phase relationships in oscillatory glycolysis [67]. In another experiment, glycogen, protein, lipid and nucleic acid concentrations were monitored in fish liver [68] using this method.
Single-molecule OMI is a very young field that holds great promise. It is not a technique, but a way of thinking that can allow us to detect individual molecular interactions, protein dynamics and signaling transduction pathways in living cells, which is difficult and sometimes impossible to achieve using conventional techniques [46]. For example, Sonnleitner et al. observed that the voltage-gated ion channel does not directly open or close under the conditions of single-molecular interaction using this approach [69]. Karymov et al. reported a method that could be used to directly follow junction branch migration in the holiday junction at the single-molecule level, and they detected that branch migration was a stepwise random process where the overall kinetics were dependent on the Mg2+ concentration [70].
Using OMI to monitor physiological processes in living cells
Monitoring gene expression and RNA localization
The fusion of gene promoter and fluorescent protein cDNA can be used to noninvasively analyze the expression pattern of a target gene, which is a very common approach used to study gene expression in vivo and is widely used in life science research. To minimize the accumulation of the background fluorescence that is generated by leaky, undegraded level (i.e. from previously expressed fluorescent protein that has not yet degraded), several chimeric EGFPs with shorter fluorescence half-lives were constructed by fusing protein [71, 72]. However, this approach leads to lower sensitivity, and cannot reveal low levels of expression. To overcome these drawbacks, a new approach that is similar to the amplification of an enzymatic reporter was applied, which was then used in “enhancer-trap” strategies [73, 74]. The principle behind this is that the fluorescence of FP, revealing targeted gene expression in different tissues, is magnified through a transcript factor, which allows weak signals to be seen.
In addition, DNA and RNA sequences and other molecules could be visualized indirectly by labeling their binding proteins with FP in vivo. For example, by using RNA binding protein MS2 labeled with GFP, the mobility of targeted RNA could be visualized in vivo [75]. In another study, a similar method was used to observe gene expression correlated with a change in chromatin structure under the progress of transcription in real time [76]. Compared with FISH in vivo, the disadvantage of this method is the introduction of a protein complex of considerable size.
Monitoring protein and subcellular organelle dynamics
FP can also be used to monitor the behavior of the targeted protein, such as its appearance, degradation, localization, translocation and interaction in vivo. For example, by using a fusion protein of GR and FP, the dynamics of the exchange of GR with its binding sequence array can be monitored [77]. In addition, visualization of the dynamics of the GFP-GR fusion protein in vivo in more physiological conditions was achieved by using GFP-GR knock-in mice [78]. Another interesting application involves identifying the localization of an unknown protein on a large scale, a method termed the “protein trap,” in which the imaging of cells containing the fusion protein of FP and a cDNA library was used to screen for target protein at the localization of interest [79].
Fusion protein that targets a given subcellular organelle is often used to study the dynamics of subcellular organelles. For example, Shaw et al. [80] revealed the dynamics of migration across the cell cortex of individual cortical microtubules in Arabidopsis by using tubulin fused to FP. In a similar study, this method was used to measure the rates of tubulin polymer growth, shortening and transition [81]. Furthermore, new subcellular structures can also be discovered by visualizing fusion FP, such as the novel discrete area in the nucleus, in which phytochrome species were induced to accumulated by light [82].
Monitoring general protein–protein interactions
Detecting the proximity of two biomolecules based on FRET is an approach widely used to observe protein interactions in real time in vivo. Most commonly, CFP and YFP are respectively fused to each of their putative interaction proteins, which are coexpressed in one living cell so that FRET between two fused FPs of interaction can be detected (Fig. 5a). A variety of protein interactions in different cells have been successfully visualized so far using intermolecular FRET, such as the interaction between phytochrome B and cryptochrome 2 [83], the rearrangement of G-protein subunits [84], and the oligomeric states of the ligands B7-1 and B7-2 [85]. Protein–protein interactions can also be imaged via protein complementation assays. The putative interaction proteins are respectively fused to two complementary fragments of one fluorescent protein, and then interaction of the proteins can reinvoke the fluorescence [46, 86], in a technique which is analogous to yeast two-hybrid assays (Fig. 6a). Using this method, Hu et al. simultaneously visualized several interactions among bZIP and Rel family transcription factors in one cell [87, 88].
Fig. 5a–dGeneral designs of FRET-based fluorescent probes. a An intermolecular probe consists of two interacting proteins that are labeled with CFP and YFP, respectively, which interact and result in FRET. b An intramolecular probe consists of CFP and YFP fused together with a cleavable linker or protein, which can be cleaved by proteolysis and disrupt FRET. c An intramolecular probe consists of sandwiching two domains between CFP and YFP, which can interact after phosphorylation or binding to calcium, resulting in a change in FRET. d An intramolecular probe consists of CFP, YFP and a protein/domain, which permits conformational change by binding to another biomolecule, leading to a change in FRETFig. 6a–dSingle FP-based fluorescent probes. a The probe consists of the fusion of two interacting proteins to two complementary fragments of one FP, respectively, which can interact and reinvoke the fluorescence. b Insertion of a conformationlly responsive domain/protein into cpYFP can lead to a change of fluorescence when its conformation is changed. c The probe consists of the fusion of two interacting proteins/domains to the amino and carboxyl termini of cpGFP, which can interact and change the cpGFP fluorescence. d By using mutagenesis, AFP can be engineered to be directly sensitive to a small molecule, such as Cl−, H+
Monitoring protease and kinase activity
The first reporter of proteolysis consists of BFP and GFP fusing together with a protease-sensitive linker, which exhibits the FRET phenomenon. Proteolysis can disrupt the FRET by separating the donor and acceptor fluorescent proteins [89]. In recent studies, fusion proteins of CFP, YFP and a specific recognition sequence were used to measure the different caspase proteolytic activities during different apoptosis pathways (Fig. 5b) [90–92]. In another study, YFP-Bid-CFP fusion protein was used to visualize the activation of Bid protein by proteolytic cleavage, and translocation of the cleaved Bid to mitochondria was observed directly [93].
Reporters for the activity of tyrosine kinases and serine/threonine kinases have been made by sandwiching a substrate peptide for the kinase of interest and a phosphoaminoacid-binding domain, such as Src-homology-2 (SH2) or 14-3-3 protein, between two FPs (Fig. 5c). Phosphorylation of the substrate peptide induces the formation of an intramolecular complex with the phosphoaminoacid-binding domain, which leads to a change in the FRET. Meanwhile, these indicators also report the opposite phosphatase activity. This generic concept has so far been adapted to create probes for the phosphorylation of the kinase of interest in different signal transduction pathways, including phosphorylation of protein kinase A (PKA) [94], activation of serine/threonine kinase Akt [95], and activation of Src kinase on the cell membrane [96].
Monitoring changes in calcium
Genetically encoded fluorescent indicators for calcium without cofactors that can target to specific intracellular locations were first constructed by Miyawaki et al. [97]. This indicator was termed “cameleon.” The cameleon consists of the tandem fusion of CFP, calmodulin, the calmodulin-binding peptide M13 and YFP. Binding of Ca2+ allows calmodulin to wrap around the M13 domain, which increases the FRET between flanking FPs (Fig. 5c). Replacing YFP with cpYFP [28], Venus [98] or EYFP [99] led to a new generation of cameleon that exhibited better spatial and temporal resolution as well as better environmental resistance. In order to measure high concentrations of Ca2+, the CaM–M13 interface of cameleon was re-engineered to reduce its affinity to Ca2+, which improved the imaging of Ca2+ in the endoplasmic reticulum [100].
Other indicators for calcium that are not based on FRET have also been reported. cpGFP or cpYFP can tolerate the insertion of another entire protein, and conformational change of the fusion cpFP can change its fluorescence. Therefore, the insertion of calmodulin into cpYFP (called “camgaroos”) (Fig. 6b) [14, 27] and the tandem fusion protein of M13, cpGFP and calmodulin (termed “G-CaMP”) (Fig. 6c) [101] have both been used as calcium indicators because a conformational change occurs in the fusion cpFPs upon calcium binding.
Monitoring changes in cyclic nucleotides
The first fluorescent sensor of cAMP consists of cAMP-dependent protein kinase (PKA) in which the catalytic subunit and the regulatory subunit are labeled with a fluorescein and a rhodamine, respectively. Binding of cAMP to the labeled regulatory subunit leads to the dissociation of the labeled catalytic subunit, which disrupts the FRET between the two fluorescent dyes [102]. Later, this system was developed into a genetic cAMP indicator with BFP and GFP replacing the fluorescent dyes [103]. The most recent cAMP indicator was constructed by sandwiching a full length Epac1 between CFP and YFP (Fig. 5d) [104].
A genetically encoded indicator for another cyclic nucleotide, cGMP, has been reported by Sato et al. [105]. This indicator consists of a tandem fusion of BFP, nondimerizing mutants of cGMP-dependent protein kinase Iα (PKG Iα) and GFP. An increase in FRET between the two FPs can be detected upon the cGMP-induced conformational change of PKG Iα. Another sensor, termed “cygnet-1” [106], was constructed to have high selectivity for cGMP over cAMP. Here, the truncation of PKG Iα consisted of residues 1–77 rather than 1–47 reported in a previous study [105], and it exhibited decreased FRET fluorescence upon the binding of cGMP.
Monitoring changes in transmembrane voltage
Measuring electrical activity in living cells with high spatial and temporal resolution is a fundamental problem in studies of excited cell information processing. To address this problem, Siegel and Isacoff [107] first constructed a novel fluorescent probe that could be used to measure transmembrane voltage in vivo, termed the fluorescent shaker or “FlaSh”, in which the modified GFP was inserted into a voltage-sensitive K+ channel so that voltage-dependent rearrangements in the K+ channel fusion protein could induce changes in the GFP fluorescence. A similar voltage sensor was generated by inserting GFP into another channel, the rat μI skeletal muscle voltage-gated Na+ channel, and was named “SPARC” [108]. In other studies, Sakai et al. [109] developed a new voltage-sensitive indicator based on FRET, in which changes in membrane voltage can lead to changes in FRET.
Monitoring changes in pH in living cells
The chromophore of a FP is surrounded by a hydrogen-bonding network within the β-barrel, which means that the FP is pH-sensitive because of direct interactions between its hydrogen-bonding network and external protons [12]. The majority of GFP, YFP and their circularly permuted constructions described in previous parts of this review exhibit sensitivity to acid (Fig. 6d), whereas the original AFP—Renilla reniformis GFP—and DsRed are insensitive to acid. Therefore, acid-sensitive variants can serve as ideal donors in pH indicators based on FRET, and DsRed is the most desirable acceptor [110]. Another pH sensor has been generated by using a new group of pH-sensitive GFP variants, termed “deGFPs,” which exhibit rapid changes in emission from green to blue as the pH decreases, and these are suitable for ratiometric measurements in vivo [111, 112].
Monitoring signal transduction from cell to cell
Scientists have also monitored intercellular changes in biomolecules and physiological events, which have been visualized in single cells in real time in a cell population. For example, Bedner et al. monitored the intercellular permeability to cAMP of six different gap junction channels by using sensors that are highly sensitive to cAMP concentration [113]. The change in calcium concentration from cell to cell was also monitored via calcium dyes [114]. In addition to these biomicromolecules, the propagation of intercellular apoptotic or survival events in monolayer cells mediated by the gap junction channel were also studied through OMI [115].
Using OMI to investigate tissue structure and function
It has recently been shown that SHIM can be used to directly detect several structural protein arrays in tissues, such as collagen arrays in mouse [116], neurons and muscular structures of the pharynx in C. elegans [117] without the need for fluorescent labeling. SHIM also readily retrieves more detailed molecular information than that obtained through the application of fluorescent labeling. Therefore, SHIM can be used to study or diagnose several diseases that are related to the assembly polarity of microtubule complexes in native brain tissue [118]. In addition, SHIM has proven to be crucial to neuroscience investigations in thick tissue preparations. For example, the fast neuronal membrane potential transient in mammalian brain slices labeled with FM4-64 dye was investigated by this method [119]. By labeling with another dye, the action potential was recorded with high temporal and spatial resolution on soma and neurite membranes [120].
OCT also has the ability to perform in situ, real-time imaging of tissue pathology, and it can be used to guide excisional biopsy in order to reduce false negatives caused by sampling errors. In particular, OCT enables the internal architectural morphology of the retina to be visualized noninvasively, and it can be used to diagnose and monitor retinal diseases, which cannot be achieved through any other methods [121]. The development of high-speed OCT imaging combined with small fiber-optic probes has enabled in vivo endoscopic imaging, such as the visualization of the oral cavity [122], the larynx and the bladder [123]. In addition, OCT can perform functional imaging of tissues, such as brain activity [124], inflammatory/neoplastic morphologic changes [125]. Raman spectroscopy (RS) is another powerful diagnostic tool that enables tissue identification and classification, as demonstrated by measurements of the brain tissue of a six-month-old pig by fiber-optic probes [126] and analysis of the molecular composition of human bronchial tissue structures [127]. Also, a combination of confocal RS and LSCM has been employed to obtain detailed information about the subsurface structures in the skin with high spatial resolution in a completely noninvasive manner [128].
OMI of organisms
Employing fluorescent protein
When performing whole-body OMI of organisms, the utilization of genetically encoded fluorescence probes has enabled scientists to decipher spatial and temporal changes in biological events inside complex organisms [129]. Over the past few decades, FPs have been used to generate various fluorescent probes that have been used to image physiological and biochemical events in the monolayer cells. Extracting information from whole-body imaging is more relevant to real physiological conditions, but up to now, only a few fluorescent indicators have been used in complex organisms to visualize physiological activity in vivo.
To study the feeding behavior of C. elegans, Kerr et al. [130] introduced a calcium indicator based on cameleon to C. elegans, which allowed them to measure the Ca2+ signals evoked in pharyngeal muscles and individual neurons under the stimulation of feeding in vivo (Fig. 7a,b). Similar visualization of cameleon localized in other tissues was used to study the functions of serotonin and G proteins [131], the role of G proteins in signal transduction [132], and the ASH neuron response [133] in C. elegans. The spatial and temporal representation of odorant-evoked Ca2+ signals in the Drosophila brain was explored using cameleon [134] and G-CaMP (Fig. 7c–f) [135]. Calcium imaging was also used in other animals, such as zebrafish (Fig. 7g–i) [136] and mouse [137, 138]. In addition, an indicator with pH sensitivity was used to visualize spatial patterns of defined neuronal activity in the mouse [139]. More recently, the proteolytic activity of calpain was visualized by introducing fluorescent indicator into living mouse muscle, which gave the first 3-D imaging of FRET in vivo [140].
Fig. 7a–kWhole-body OMI using FPs and QDs. Imaging of Ca2+ signals in pharyngeal muscles under the conditions of noncontraction (a) and contraction (b) (red color indicates higher calcium) in transgenic C. elegans expressing cameleon. Reproduced from [130] with permission. c–f Ca2+ signals evoked by different odors in Drosophila brain expressing G-CaMP. Reproduced from [135] with permission. Imaging of a three-day-old transgenic fish (g) carrying the cameleon and its RB neurons (h) by confocal optical section, and the change in the fluorescence ratio (i) (representing the calcium concentration) in an RB neuron under electrical stimulation of the skin. Reproduced from [136] with permission. j Simultaneous multicolor imaging in a mouse injected with QD-encoded microbeads; k QD imaging of a prostate tumor in the mouse. Reproduced from [33] with permission
Employing QDs
Due to several advantages of QDs over FP, QDs have been used to image large animals in vivo, which holds great promise for clinical applications, especially in the imaging of tumors. Akerman et al. [141] report the application of QDs coated with specific targeting peptides to the imaging of different tissues in tumor in vitro, but this QD probe cannot be detected in living animals. Gao et al. [33] report on a new class of multifunctional QDs that were encapsulated with an ABC triblock copolymer and then linked with a tumor-targeting antibody. Using subcutaneous injection or systemic injection of these QDs into the mouse, simultaneous multicolor fluorescence imaging of the prostate tumor with efficient background removal and precise delineation of weak spectral signatures based on wavelength resolution was achieved (Fig. 7j,k). More recent research showed that the utilization of QDs in combination with GFP labeling can differentiate tumor vessels from both perivascular cells and the matrix, and QDs linked to bone marrow-derived precursor cells can visualize tumor vasculature [142]. These OMI techniques, as well as their applications at different biological levels, are summarized in Table 1.
Table 1OMI in systems biologyTechniquesPhysical processesResolutionSensitivityPenetrationObservation methodsProbesIndicator localizationReferencesWFM LSCM MPLSMRNA transcription and expressionHighHighHighDFI, FRETFPsIn vitro, cells, tissues, organism[63, 64, 71–76]Protein localization and dynamicsDFIFPsIn vitro, cells, tissues, organism[51, 52, 65, 77–79]Subcellular organelle dynamicsDFI, FRAPFPsCells[80–82]Protein interactionFRET, FLIM, FCSFPs, cpFPsCells, mouse[46, 47, 49, 57, 59, 60, 83–88, 140]Protease and kinase activityFRETFPs[89–96]CalciumDFI, FRETFPs, cpFPsCells, C. elegans, fish, mouse, Drosophila[14, 27, 28, 97–101, 130–138]cAMP, cGMPFRETFPsCells[102–106]Electrical activityDFI, FRETFPsCells[107–109]pHDFI, FRETFPsCells, mouse[110–112, 139]TumorDFIQDsMouse[33, 141, 142]SHIMBiomacromoleculesLowHighLowNI–Mouse, C. elegans, brain tissue[116–118]Membrane potentialNIDyesCells, brain tissue[119, 120]OCTMorphology of tissueHighLowHighEI–Human[122, 123]Activity and disease of organEI–Human[124, 125]RSComposition of structureLowHighLowNI–Brain tissues, bronchial tissue[126, 127]FT-IRKinetics of reaction intermediatesLowHighLowNI–Yeast extract, fish liver[67, 68]Abbreviations: DFI, Direct fluorescent imaging; NI, nonlinear imaging; EI, light-echo imaging
Future development
As described above, OMI is a feasible analytical technique for monitoring the dynamics of biological events in vivo and in real time anywhere from single-molecule to whole-body level, which makes it well-suited to systems biology measurements. Its noninvasive, ultrasensitive, high-resolution and real time nature has resulted in the widespread use of OMI. However, its disadvantages are also very apparent, such as its high cost, its low throughput, and its limited detectable depth, which is a particular hindrance in whole-body analysis. It should be pointed out that OMI is still far from fulfilling all of the requirements of systems biology (as concluded in our previous paper [8])—in terms of sensitivity, selectivity or specificity, linear range of quantitation, throughput, robustness, flexibility and cost—that are definitely required in genomics, transcriptomics, proteomics, and metabolomic profiling and dynamics, although it it clear that OMI is a promising method for these types of bioanalysis.
In the near future, several routes to improving OMI for systems biology need to be followed. Firstly, it is crucial to develop better molecular probes for OMI; in other words probes that have improved biocompatibilities and high quantum yields, are easy-to-tag, display multiple colors and have better penetration capabilities. The importance of the molecular probe to OMI cannot be overstressed. Current endeavors in QD development are heading in a good direction but this field is still in its infancy [143]. However, this doesn’t mean that the development of small organic dyestuffs and biogenetic FP is less important: on the contrary, applications of these two kinds of molecular probes will still constitute most OMI studies [144]. Secondly, the integration of several modalities or approaches should be emphasized, since it makes OMI much more informative. Thirdly, efforts to improve OMI instrumentation are also important, since they will result in better time and space resolution and better sensitivity. For example, the 4Pi technique has improved the spatial resolution of OMI down to the mid-nanometer level, which is over the diffraction limit. The rapid progress currently being achieved in nanotechnology and nano/microelectromechanical systems looks set to lead to the manufacture of low-cost and high-efficiency OMI instruments [145]. Finally, the application of OMI to the life sciences is important; it is the reason behind the development of OMI. In particular, the application of OMI to systematic measurements of biological systems will be of great interest, which is also the aim of systems biology.

optical molecular imaging
systems biology
review
quantum dots
fluorescent protein