Next Article in Journal
Tellurium-Doped, Mesoporous Carbon Nanomaterials as Transparent Metal-Free Counter Electrodes for High-Performance Bifacial Dye-Sensitized Solar Cells
Next Article in Special Issue
From Single-Core Nanoparticles in Ferrofluids to Multi-Core Magnetic Nanocomposites: Assembly Strategies, Structure, and Magnetic Behavior
Previous Article in Journal
Wide-Angle Polarization-Independent Ultra-Broadband Absorber from Visible to Infrared
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Nanomaterials
Volume 10
Issue 1
10.3390/nano10010028
nanomaterials-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Luminophore and Magnetic Multicore Nanoassemblies for Dual-Mode MRI and Fluorescence Imaging

by
Lénaïc Lartigue
*,
Marina Coupeau
and
Mélanie Lesault
Université de Nantes, CNRS, CEISAM UMR 6230, F-44000 Nantes, France
*
Author to whom correspondence should be addressed.
Nanomaterials 2020, 10(1), 28; https://doi.org/10.3390/nano10010028
Submission received: 3 December 2019 / Revised: 15 December 2019 / Accepted: 17 December 2019 / Published: 20 December 2019
(This article belongs to the Special Issue Multicore Magnetic Nanoparticles for Biomedical Applications)
Browse Figures
Versions Notes

Abstract

:
Nanoassemblies encompass a large variety of systems (organic, crystalline, amorphous and porous). The nanometric size enables these systems to interact with biological entities and cellular organelles of similar dimensions (proteins, cells, …). Over the past 20 years, the exploitation of their singular properties as contrast agents has led to the improvement of medical imaging. The use of nanoprobes also allows the combination of several active units within the same nanostructure, paving the way to multi-imaging. Thus, the nano-object provides various additional information which helps simplify the number of clinical procedures required. In this review, we are interested in the combination between fluorescent units and magnetic nanoparticles to perform dual-mode magnetic resonance imaging (MRI) and fluorescent imaging. The effect of magnetic interaction in multicore iron oxide nanoparticles on the MRI contrast agent properties is highlighted.

1. Introduction

Medical imaging allows for non-invasive anatomical or functional diagnostics to detect dysfunctions or specific diseases. Physical principles of the considered technique determine the spatial resolution, the sensitivity level and the type of tissues or biological phenomena to be detected [1]. Among the most commonly used in clinics, we can mention X-ray computed tomography (CT scan), magnetic resonance imaging (MRI), ultrasound imaging, and nuclear imaging (single photon emission computed tomography, SPECT or positron emission tomography, PET) [1,2]. For all imaging techniques, contrast agents (i.e., active units) are generally essential and the use of nano-objects suggests promising perspectives [3]. Indeed, nano-objects enable the combination of different active units inside the same system. Thus, the development of multimodal nanoprobes made of various imaging units can advantageously multiplex images to avoid false-positive results, thereby providing earlier and more reliable diagnosis. Among the different possible combinations, growing interest has been shown in nanosystems comprising magnetic nanoparticles (MNPs) for MRI and luminescent entities for fluorescent imaging in recent years (Figure 1) [4]. Luminescence imaging, thanks to its multiple-label possibility, its high sensitivity and spatial resolution, is widely used to follow biological processes or in histopathology. However, extinction phenomena (diffusion and absorption of light by tissue) limit the depth penetration of this imaging technique. By contrast, MR present an unlimited depth penetration and greater soft-tissue contrast. Nevertherless, the low sensitivity of the MR technique makes it difficult to distinguish benign from malignant disease even with long acquisition time. So, combining these two safety techniques (using non-ionizing radiation) allows advantage to be taken of the high sensitivity and spatial resolution of luminescence imaging, associated to the good spatial resolution and deep tissue penetration of MRI. This combination is particularly interesting for correlating in vitro monitoring and in vivo tracking [5].
This review proposes to compare the different approaches envisaged to design dual-mode nanomaterials. A special emphasis is made on nanosystems embedding multicore magnetic nanoparticles. First, descriptions of the techniques and the different functional units are presented. Then, effects of self-assemblies magnetic iron oxide nanoparticles are discussed. Finally, the strengths and weaknesses are assessed of the three methods leading to magneto-fluorescent nanosystems: (1) association by covalent bonding; (2) encapsulation in matrices; (3) dispersion in nanoassemblies.

2. Active Units

2.1. Magnetic Entities for Magnetic Resonance Imaging (MRI)

MRI is a non-invasive technique using radio frequency (RF) pulses and strong magnetic fields to create images of internal organs and structures. MRI is based on the nuclear magnetic resonance of the hydrogen nuclei (proton) mainly those of water, which is abundant in most organs. The relaxation mechanism of these protons is explained in Figure 2A.
Hydrogen atoms have only one nuclear spin that induces a nuclear magnetic moment. In the presence of an external static magnetic field B0, the spin magnetic moments of protons, represented by a total magnetic moment, denoted M0, will precess around the direction of the field B0. A very short perpendicular RF B1 pulse tilts the magnetic moment of protons in (Oz) plane. Their relaxation to the initial state is measured after the pulse. Return to the equilibrium position (i.e., M0 in the same direction as B0) can be decomposed into two components Mz and Mxy (Figure 2B). Mz, the component along the axis (Oz), which is associated with the longitudinal relaxation R1 and the relaxation time T1 (R1 = 1/T1). The Mxy component in the plane (Oxy) is related to the transverse relaxation R2 and the relaxation time T2 (R2 = 1/T2). The longitudinal relaxation R1 corresponds to a spin-lattice relaxation, i.e., exchange energy of the spins with the external medium [6]. The transverse relaxation R2 corresponds to a spin-spin relaxation due to interactions between spin-magnetic dipole moments (Figure 3A). This transverse relaxivity is also sensitive to local magnetic field heterogeneities induced by species (tissues or materials placed within the field) possessing different magnetic susceptibility [6]. This local inhomogeneities produce, in addition to spin-spin relaxation, an additional dephasing fields. Both together induce an effective transverse relaxation called T2* and defined as:
1 T 2 * = 1 T 2 + γ Δ B i n h o m .
where γ is the gyromagnetic ration and ΔBinhom the local magnetic field inhomogeneity.
In practice, the T2* relaxations are observed in gradient-echo (GRE) sequences by contrast the spin-echo sequence use a 180° rephasing to avoid the effect of field inhomogeneities and record the T2 relaxation.
This technique allows the temporal or spatial (2D or 3D) reconstruction of living tissues, rich in protons, with an excellent spatial resolution, no limit in depth and without the use of ionizing radiation. The difference in image contrasts in MRI is defined by either intrinsic tissues properties (e.g., inhomogeneity in viscosity or water concentration) or operator-selected pulse sequence parameters (repetition time, TR, or echo time, TE). However, the MRI has important limitations, including a long acquisition time and the impossibility of achieving large areas in a single acquisition images (whole body for example). Another weak point of this technique is its low sensitivity, sometimes making difficult the distinction of pathological tissues. Thus, paramagnetic agents (gadolinium complexes, positive contrast) or superparamagnetic agents (iron oxide, negative contrast) are commonly used to enhance the native contrast between different tissues, which makes diagnosis more reliable. Contrast agents (CAs) dramatically shorten the T1 and T2 of water and their presence is easily detected in MRI images at levels as low as 0.1 mmol L−1. The efficiency of MRI contrast agents is determined by measuring the nuclear relaxivities r1,2 defined by Equation (2):
r i = [ ( 1 T i ) m e a s . ] − ( 1 T i ) d i a . c
where (1/Ti)meas is the value measured for the sample with concentration c (mmol L−1) of magnetic center, and (1/Ti)dia refers to the nuclear relaxation rate of the diamagnetic host solution.
The first MRI T1-contrast agent, Magnevist® (Bayer Schering Pharma AG, Lerverkusen, Germany), approved in 1988 is a gadolinium-based positive contrast agent stabilized with diethylenetriaminepentaacetic acid (DTPA). Even now, most of the MRI agents on the market are gadolinium-based complexes. If Magnevist® remains the most used with 51% of market share, gadolinium-based complexes with different polyaminocarboxylate ligands have been developed: Omniscan® (25% market share, GE Healthcare, Chicago, IL, United States) stabilized with the linear DTPA-BMA (diethylenetriamine pentaacetate bismethylamide) ligand and Dotarem® (12%, Guerbet SA, Villepinte, France) with the macrocyclic DOTA (1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid) ligand [7]. However, over the past 10 years, studies have shown that gadolinium ions are released (i) by transmetallation when chelated by linear complexes (competitively with other endogenous ions such as Zn2+), or (ii) by acid catalyzed dissociation (even at physiological pH) when associated by macrocycles [8]. Moreover, biological ligands, such as adenosine triphosphate, block their activity [8]. Gd3+ ions are excreted via the renal system and therefore contraindicated for patients with renal insufficiency [9]. Therefore, since 2017, Magnevist® and Omniscan®, in particular, have been recommended for withdrawal in Europe and the US market [10,11]. In parallel, superparamagnetic nanoparticles have been developed as a contrast agent for T2-weighted imaging. The efficiency of magnetic NPs is characterized by the largest possible value of the transverse relaxivity, r2, and r2/r1 ratio. Iron oxide nanoparticles (magnetite, Fe3O4 and maghemite, γ-Fe2O3) are the most commonly used T2-contrast agents due to their low toxicity, remarkable magnetic properties (high magnetic susceptibility and saturation magnetization) and their great stability in biological environments [12]. As described in Table 1, negative contrast agents have been developed in the past but have since been withdrawn for commercial reasons. These contrast agents are multicore nanoassemblies of 3–5 nm magnetic nanoparticles coated with a polymer-based ligand (dextran, carboxydextran, polystyrene). One of the major shortcomings of these commercial agents was the relatively poor improvement in the contrast of imaging (r2 and r2/r1 ratio does not exceed 200 s−1 mM−1 and 20 respectively).
To improve the effectiveness of MRI negative contrast agents, modification of their size, shape, state of assembly and surface functionalization was needed [15,16]. In particular, the use of multi-core nanoassemblies with larger spherical magnetic nanoparticles [17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43] has been envisaged (Table 2). Multi-core nanoparticles consist in self-assembled magnetic nanoparticles (magNAs) providing a very high effective magnetic moment per magNAs [44,45,46,47,48]. The effective magnetic moment, μeff, is dependent on the number of nanoparticles per assembly N, the volume saturation magnetization MS (in A m−1), and the assemblies’ diameter which can, in first approximation, be assimilated to the hydrodynamic diameter dH [44,45]:
μ e f f = N 1 6 π d H 3 M S
According to the diameter dH of the contrast agent, three regimes are successively achieved (Figure 3B): first the motional average regime (MAR), then the static dephasing regime (SDR) and finally the echo-limiting regime (ELR) [19,28,49,50]. Vuong et al. [50] defined the transition from the MAR to SDR as a function of ΔωτD factor. The angular frequency shift experienced by a proton at the equator of the particle, Δω is calculated following Equation (4).
Δ ω   =   γ μ 0 M S 3
and the translational diffusion time of the proton in the magnetic field inhomogeneities, τD is deduced from Equation (5):
τ D = d H 2 4 D
where the gyromagnetic factor of the proton, γ = 2.67513 × 108 rad s−1 T−1; the magnetic permeability of vacuum, μ0 = 4π × 107 T m A−1; and the water translational diffusion time, D = 3.10−9 m2 s−1 at 37 °C.
If ΔωτD < 1, the r2 relaxivity is described by the MAR and is expressed by Equation (6):
r 2 = 4 γ 2 d H 2 ( μ 0 M S ) 2 ν m a t 405 D
where the iron oxide molar volume, νmat = 1.57 × 10−5 and 1.50 × 10−5 m3 mol−1 for maghemite and magnetite respectively [50].
If ΔωτD > 1, the r2 relaxivity followed Equation (7) of the SDR:
r 2 * =   2 π γ ( μ 0 M S ) ν m a t 9 3 ≈   r 2
where r2* is the apparent transverse relaxivity which in addition of the r2 takes into account of the local field inhomogeneity.
In MAR, the relaxivity is a function of dH2 and increases with the outer sphere diameter assimilated to dH. In SDR, the relaxivity has reached a maximum and is independent of the outer sphere diameter [28,49]. Finally, when the dH increases too much, transverse relaxivity is described by ELR. In this regime, the transverse relaxivity decreases when the size of the assemblies increases. As described in Figure 4, self-assembly of magnetic nanoparticles into silica matrix [20,27], micelle [24,25,28,30,32,34,35,38,42,43], polymersomes [17,18,30,31,36,37,39,41], or liposomes [22,26,29,40] are synthetized. Depending on the structure type, the incorporated MNPs should be hydrophilic or hydrophobic. Their dispersions can be done in two ways: in the shell (structure noted type I) or in the core (structure noted IIa-d). The MNPs used are prepared by two synthetic routes: coprecipitation (CP) or thermal decomposition (TD). The co-precipitation method is based on the hydrolysis of transition metal ions in aqueous solution [51,52]. The reaction takes place in alkaline aqueous solution with an optimal pH around 8.5–10 [53]. The nanoparticles obtained present a very broad size distribution and a size sorting is necessary [54]. The stability of these nanoparticles is insured by the surface charge: positive and in the form −OH2+ for pH between 1 and 3.5; or negative and in the form of −O− for pH 9–11 [55,56]. The easy grafting of phosphoric, phosphonic or carboxylic acid derivatives hydrophilic (citrate [57], polyacrylic acid [58,59], …) or hydrophobic (oleic acid [54,56], Beycostat NB09 [18]) on the MNPs surface, allows respectively their transfer in neutral aqueous solution or organic solvent. By this method, depending on the base used and their size, the MNPs have a saturation magnetization of around 50–70 A m2 kg−1 [52,53,60] which can be increased close to the bulk material value (80 A m2 kg−1) after an hydrothermal treatment [61].
An alternative approach to the synthesis of monodisperse iron oxide nanoparticles is high-temperature organic phase decomposition of an iron precursor [62,63,64,65,66,67]. This reaction involves the thermal decomposition of a preformed organometallic precursor dissolved in a high-boiling organic solvent (usually refluxed) in the presence of capping ligands such as long-chain alkyl surfactants. The most commonly couple capping ligand/precursor used is oleic acid/iron(III) acetylacetonate. The thermal decomposition method yields very narrow particle-size distributions (standard deviation between 0.10 and 0.15) as well as excellent crystallinity and shape control. By this method, the hydrophobic MNPs possess relatively high saturation magnetization around 70–85 A m2 kg−1 [68,69,70]. Recently, several groups have shown the possibility to prepare multi-core nanoparticles without an organic matrix by a polyol route [71,72,73]. In this route the high-boiling polyol compound acts as both solvent and reductant agent of the iron precursor, an iron chloride salt. At high temperature, multicore nanoparticles with saturation magnetization in the 60–80 A m2 kg−1 range, and with different size are obtained by adjusting the used polyol (ethylene glycol, or mixture of diethylene glycol with N-methyldiethanolamine or sodium hydroxide) and the experimental condition [71,73,74]. Thanks to the increase of the magnetic moment per assembly, multicore MNPs provide very high value of r2 up to 1400 mM−1 s−1 (Figure 4) [28]. Moreover, the clustering of MNPs significantly reduces the exchange surface between magnetic materials and water molecules inducing a decrease in the r1 value. Thus, these two phenomena lead to very good r2/r1 ratio (up to 2000) [35]. Significant values of r2 (>500 mM−1 s−1) are obtained regardless of the size of the MNP used (from 6 to 16 nm), the type of organization, or the chemical route (CP, TD, polyol) adopted to prepare the magnetic nanoparticles. This observation clearly demonstrates the primacy of the total magnetic moment of assembly over the nanoparticle alone [20,26,30,31,36,38,39,41,49,74,75]. Optimal dH values are in a wide range between 70 and 250 nm depending on the systems considered. However, it is noted that this optimum is a function of the individual size of the MNP used and shifts towards low dH values when the size of the MNPs increases [28,32,35,38]. The weight percentage of iron oxide nanoparticle in the assembly (wt% IO) is also an important parameter. An increase of the value of r2 and r2/r1 ratio is observed when wt% IO increases [18,29,30].

2.2. Nanoparticles Composed of Luminophore for In Vivo Fluorescence Imaging

Fluorescence microscopy consists of visualizing emitted photons by tissues (autofluorescence) or by exogenous luminophores—after an excitation of higher energy. The use of exogenous luminophores allows the acquisition of specific physiological tissue and cell imagery, quickly and without ionizing radiation [76,77,78]. In this way, cells activity is investigated at different levels: investigation of Ca2+-pathway [79], or tracking cellular metabolites [80], biological macromolecules [81] as proteins [82]. The sensitivity of the technique is related to the specificity of the luminophore: its quantum yield and its brightness [83]. The fluorescence quantum yield, Φ, is defined as the number of emitted photons per the number of absorbed photons. Its calculation requires a reference as described below [84]:
ϕ x = ϕ r e f ·   I x I r e f · A r e f A x ·   n x 2 n r e f 2
where Φref, the known fluorescence quantum yield of a reference luminophore; Ix and Iref are the integral of the corrected emission signal at the same excitation wavelength of the luminophore and the reference, respectively. Ax and Aref are the absorbance values at the excitation wavelength of the luminophore and the reference, respectively. nx and nref the solvant refractive index of the luminophore and the reference respectively.
The brightness, B, is defined as the product of the molar absorption coefficient at the excitation wavelength, ε(λex.), and the fluorescent quantum yield, Φ:
B =   ϕ x   ·   ε ( λ e x . )
This value is the analytical parameter defining the sensitivity of the luminophore. A large panel of luminescent compounds are developed including, fluorescent molecules, fluorescent proteins, polymer dots (Pdots) or quantum dots (Qdots) [83,85,86,87]. Small molecules such as BODIPY, rhodamine or fluorescein derivatives (Figure 5A), are easily accessible, biocompatible and present tunable emissive properties from 200 to 800 nm by chemical derivation [88,89]. Moreover, these derivatives can be modified to probe organelles or cellular membranes [90]. Thus, mitochondria or lysosomal compartments are stained respectively by Rhodamine 123 (λmaxem = 529 nm) [91] or LysoTracker® Red (λmaxem = 590 nm) [92]. Nucleus staining is achieved by fluorophore binding to the base pairs of double-stranded DNA without pair specificity for propidium iodide dye or with a specificity of A-T regions for 4′,6-diamidino-2-phenylindole (DAPI) and Hoechst 33342 dye [93]. The blue (λmaxem = 450–490 nm) labeling of the nuclear DNA can be conducted by DAPI for fixed cells or Hoechst 33342 for live cells [93]. Propidium iodide (λmaxem = 617 nm) stains preferentially the nucleus of permeable cells and is used as necrotic label. These systems generally have a relatively high brightness around 104–105 L mol−1 cm−1. Nanoprecipitation of hydrophobic fluorophore gives fluorescent organic nanoparticles (FON) [94,95,96,97,98,99]. These systems are composed of a multifold of dyes (104 to 105) that are neither covalently linked nor diluted in an inert matrix, which yields highly bright structures (106–107 M−1 cm−1) [99,100]. Under mono- or biphotonic excitation, in cellulo imaging showed that these nanoarchitectures appeared as bright as quantum dots, allowing their use for tracing cancer cells and macrophages [101]. Fluorescent proteins have similar brightness of non-assembled fluorophore and allow better biocompatibility [102]. Among these proteins, the green fluorescent protein (GFP) is the more studied [103,104,105,106,107]. This green protein (λmaxem = 510 nm) has been isolated in 1960 from the jellyfish, Aequorea victoria. The discovery and development of GFP led O. Shimomura, M. Chalfie and R. Y. Tsien to the 2008 Nobel Prize. However, it has low photostability and a high cost of production [108]. Pdots are appreciated for their brightness properties (105–106 L mol−1 cm−1) and their low cytotoxicity [109]. They are conjugated polymers based on aromatic rings (Figure 5B) as poly[2,5-di(3,7-dimethyloctyl) phenylene-1,4-ethynylene] (PPE), poly[2-methoxy-5-(2-ethylhexyloxy)-1,4-phenylenevinylene] (MEH-PPV), polyfluorene derivitative as poly(9,9-dioctylfluorenyl-2,7-diyl), PFO (λmaxem = 435 nm); poly[(9,9-dioctyl-2,7-divinylene-fluorenylene)-alt-co-(2-methoxy-5-(2-ethylhexyloxy)-1,4-phenylene)], PFPV (λmaxem = 510 nm); poly[(9,9-dioctylfluorenyl-2,7-diyl)-co-(1,4-benzo-{2,1′,3}-thiadiazole)], PFBT (λmaxem = 545 nm) [110,111,112,113,114,115,116,117]. By controlling the operating parameters, self-assembly of these polymers gives nanoparticles, (typically from 10 to 50 nm) [99,110,111,112,113,114,115,116,117]. Nevertheless, a decrease in quantum yield of nearly 70% is noted when the size of a self-assembly of polybutylene terephthalate (PBT) goes from 10 to 40 nm [110,111,112,113,114,115,116,117]. Finally, Qdots are semi-conducting nanocrystals with high brightness (106 L mol−1 cm−1), high photostability and narrow emission signal [118,119,120]. As described in Figure 5C, the emission wavelength depends on the size of the crystal, and can vary from blue (380 to 440 nm) for smaller Qdots (~2 nm diameter) to red (605 to 630 nm) for larger particles (~5 nm diameter) [121]. Actually, the core shell CdSe/ZnS Qdots are the most common with a 90% luminescence quantum yield [122,123]. Yet, toxicity problems due to the presence of heavy metals and photoblinking are generally noted although efforts have recently been made to avoid these pitfalls [124,125,126].
Moreover, its application for in vivo studies has been limited for a long time because of (i) tissue autofluorescence, which decreases the signal-to-noise ratio (and therefore sensitivity), (ii) the absorption of photons by the biological tissues, which respectively induce an important depth limit. To overcome this problem, fluorophores emitting in the near infra-red have been developed. Indeed, in this area, the overall absorption of biological tissues is minimal: it is called the first biological window (700–950 nm) [127]. Thus, near infra-red fluorescence imaging (NIRF) faces limited light penetration into biological tissues, is used in preclinical small rodents studies, and has revealed a very promising potential for guided surgery [128,129]. If the association of two active units within the same system is very promising for multimodal imaging, precautions have to be taken to preserve the properties of each unit. This is especially true for fluorescent systems where magnetic nanoparticles units can deactivate or absorb part of the emitted light. The way to assemble these units is therefore a crucial parameter.

3. Magneto-Fluorescent Nanosystems

3.1. Association by Covalent Bonding (Nanoparticles)

Among the different association methods, grafting fluorescent entities on the surface of magnetic nanoparticles represents a simple approach (Figure 6) [130,131,132,133]. For example, fluorescent dyes (rhodamine B, λmaxem = 578 nm or fluorescein derivatives, λmaxem = 516 nm) are coupled to iron oxide surface [130]. These nanoparticles allow in cellulo motions of endosomes to be followed when exposed to a magnetic field gradient. Near-infrared (NIR) dyes have been also grafted as IR-820 cyanine derivative (λmaxem = 900 nm) [132] or dialkyl carbocyanine (λmaxem = 780 nm) [133] to obtain fluorescent systems emitting in the first biological window. Iron oxide multicore nanoparticles assembled by hydrophilic polymer have been envisaged to improve the MRI contrast agent properties [131,133]. Thus, the use of multicore Ferahme® iron oxide nanoparticles (dcore = 6–7 nm, dH = 17–31 nm) coupled to TO-PRO®-1 (λmaxem = 531 nm) gives bifunctional nanoparticles with a transverse relaxivity, r2 = 122 mM−1 s−1 (0.47 T, r2/r1 = 5) [131]. This transverse relaxivity is still improved using 8 nm iron oxide nanoparticles (r2 = 202 mM−1 s−1, r2/r1 = 3.8 at 0.47 T) embedded in polyacrylic acid matrix (dH = 90 nm) [133]. Moreover, these systems present NIR-emissive dye using a dialkylcarbocyanine as fluorophores (λmaxem in the region 751/780 nm and ε > 125,000 cm−1 M−1). The association of QDots, known to be brighter than small molecules, with magnetic nanoparticles is also envisaged [134,135,136,137,138,139,140]. The most common approach used is to prepare core–satellite systems. In this approach the core is composed of iron oxide nanoparticles surrounded by quantum dots (usually CdSe/ZnS) [134,135,136,140]. Pahari et al. have recently described an invert strategy where quantum dots (3.2 nm CdSe nanoparticles) are in the core and a shell of iron oxide is growned around (thickness of 1.3 nm) [139]. By this approach, a very good transverse relaxivity (r2 = 304 mM−1 s−1 at 9.4 T) is noted. Although these systems are easily synthetized, the close proximity between the luminophores and the metal core leads to a strong emission quenching. Indeed, electronic energy or electron transfers between both entities can take place while iron oxide nanoparticles significantly absorb at wavelengths less than 450 nm. The choice of fluorophores and their distance from the metallic core will therefore be essential. Moreover, direct exposure of fluorophores to the surrounding environment can modify the emissive properties of the system. Finally, the requirement of high colloidal stability of the final nanoassembly in aqueous solution excludes extensive grafting of fluorescent entities, especially if the latter are organic and hydrophobic. All together, these limitations produce low-emissive imaging agents.

3.2. Encapsulation in Silica Matrix (Nanostructure)

To protect the luminophores from quenching by the surrounding medium, the encapsulation of magnetic nanoparticles (γ-Fe2O3) and fluorescent units (small molecules, e.g., rhodamine or FITC derivatives; [141,142,143,144,145] or Qdots like CdSe/CdZn, CdS or CdZnS [146,147,148,149,150]) in mesoporous silica matrices has been envisaged (Figure 6). The encapsulation of magnetic nanoparticles and quantum dots leads to interactions between these two active units. This interaction induces (i) an increase of the magnetic anisotropy, (ii) a blue-shift of the fluorescence emission and (iii) a decrease of the quantum yield [146,149]. Silica-doped with organic dye core surrounded by magnetic nanoparticles (core-satellite assemblies) are also envisaged [141,142]. In these structures, the combination of several magnetic nanoparticles has the effect of drastically increasing the r2 value in comparison of magnetic nanoparticles alone. Lee et al. describe an r2 increase from 26.8 to 76.2 mM−1 s−1 (1.5 T) [141], and another study shows a rise from 116 to 397 mM−1 s−1 (9.4 T) [142]. These silica matrices show low cytotoxicity but provide only small amounts of encapsulated active units. In addition, although the fluorescent entities are protected from the external environment by encapsulation, they can diffuse freely outside the porous matrix as they are not covalently attached. To counter this phenomenon, hydrophobic fluorescent units amenable to self-assemble have been proposed to impart the magneto-fluorescent nanosystems with better structural stability and reduced dye leakage.

3.3. Dispersion in Nanoassemblies (Supraparticles)

The use of magneto-fluorescent nanoassemblies provides generally biodegradable systems that advantageously avoid bioaccumulation. In this context, several molecular matrices are envisaged composed of polymers [151,152,153,154,155,156,157,158,159], lipids [160,161,162,163], PDots [164,165,166,167] or organic molecule [168,169,170,171,172] (Figure 6). In this type of organization we will distinguish the assemblies with inert matrices of those composed of active units. Inert matrices are mainly composed of lipids or polymers. For instance, magnetic nanoparticles can be encapsulated in liposomes, and subsequently functionalized by a fluorescent molecule, here rhodamine [160]. The magnetofluorescent liposome exhibits good T2-contrast agent properties with r2 = 268 mM−1 s−1 at 4.7 T (r2/r1 = 85). Another approach is based on the use of polymers to combine magnetic nanoparticles and fluorescent entities. In the work of R. K. Prud’homme et al., polyethylene glycol has also been used to assemble hydrophobic NIR fluorophores (λmaxem = 800 nm), tris-(porphyrinate) zinc (II), and magnetic nanoparticles [153]. The authors show an increase in r2 from 66 to 533 mM−1 s−1 as the wt% IO increases from 4 to 16% [153]. Bawendi et al. describe the association of quantum dots and densely packed magnetic nanoparticles into “supernanoparticles” thanks to poly(vinylpyrrolidone) (PVP) ethylene glycol (EG) [152]. These assemblies (dH = 120 nm) display a high r2 value of 402.7 mM−1 s−1 at 9.4 T [152]. In the three last cases, the effectiveness of these multimodal structures has been demonstrated in murine models. Correlative treatments of the MRI and fluorescence signals have proved the preservation of the in vivo integrity of the nanoassemblies and validate the design of multimodal nanostructures.
The second type of self-assembled systems implies functional units as molecular bricks (Figure 6), thus limiting the number of organic species administrated in vivo. Hyeon et al. have assembled magnetic nanoparticles with a polyethylene glycol block polymer [157]. This polymer is functionalized with an imidazole derivative and fluorescent porphyrins (chlorin e6) whose emission is deactivated upon dye self-assembling [157]. The imidazole derivative is a pH-sensitive group which allows the disintegration of the nanostructure in the tumor medium (acidic pH) and leading to the reappearance of fluorescence. In this system (dH = 70 nm), the self-assembly of 3 nm iron oxide nanoparticles provides a transverse relaxivity r2 = 44 mM−1 s−1 at 1.5 T (r2/r1 = 13.3). Nanoassemblies incorporating Pdots and magnetic nanoparticles into phospholipid micelles improve relaxivity properties (r2 = 152mM−1 s−1 at 3.0 T) and enhance MRI contrast efficiency thereof [167]. Moreover, for these systems, important brightness and photostability under irradiation of the fluorophores have been demonstrated during in cellulo fluorescence microscopy [167]. The use of small hydrophobic molecules with iron oxide nanoparticles chelating functions is another effective approach to obtain magnetofluorescent systems [168,169,170,171,172]. In these systems, the fluorescent core composed of 105 dyes is surrounded by magnetic nanoparticles shell [168]. This architecture deals with a very effective dual-mode contrast agent with a brightness around 107 L mol−1 cm−1 and transverse relaxivity r2 = 238 mM−1 s−1 (0.47 T) [168]. This contrast agent displays excellent properties in liver imaging on small rodents both as a cellular label and as in vivo follow-up [168]. In cellulo stability of these systems could be controlled by varying stabilizing ligands [170]. The use of polyacrylic acid allows a very cohesive architecture, when the stabilization by citrate ions allows a dissociation [170]. These systems have been functionalized with polyethylene glycol-based copolymers to increase their circulation time [172]. Moreover, it has been shown that the presence of a hydrophobic tail in the copolymer increases the r2/r1 two times compared to those which are without one [172].

4. Conclusions and Future Outlook

The combination of magnetic and fluorescent units into a single nanomaterial provides imaging agents from in cellulo (fluorescence imaging) to in vivo (MRI) experiments through the imaging of small rodents (colocalization of fluorescence and MR signal). These very promising nano-objects must be carefully synthesized to preserve the physical properties of each active unit. We have assessed three approaches to address the issue: (i) association by covalent bonding, (ii) encapsulation of matrix, (iii) dispersion in nanoassemblies. The three systems allow multicore magnetic nanoparticles to be obtained. This configuration allows to obtain high transverse relaxivity value (r2 > 200 mM−1.s−1). The first and simplest method provides nevertheless magneto-fluorescent systems with low brightness. The second significantly improves the brightness but produces systems that are not stable over time. Finally, the third method seems to be, at the moment, the most promising because it provides ultra-bright, high MRI sensitive and stable nanoassemblies allowing a long-term follow-up. Thanks to these structures, the encapsulation of drugs can also be envisaged. Indeed, drug delivery by supramolecular nanoassemblies (supraparticles) allows to associate a wide diversity of active units simply assembled by weak bonding. These systems offer combinatorial modularity, tunable properties and biodegradability. Thus, supramolecular nanoassemblies appear as very promising drug carriers able to vectorize a large amount of drug, monitor its delivery thanks to imaging agents, and control its release with remote tunable stimuli.
In order to transfer these promising multimodal nano-objects from the laboratory to the clinic, a number of bottlenecks still need to be addressed. The in vivo biodistribution and fate of nano-systems remain the main obstacles. These points are crucial for any object designed for nanomedicine but are still more complicated in the case of a heterogeneous object and in which each unit is capable of behaving differently in a biological environment. Thus, for these complex structures it is necessary to ensure their in vivo integrity during the imaging procedures. In addition, it is necessary to know the long-term fate of all the active units i.e., their possible bioaccumulation or long-term toxicity. Thus, a back and forth between chemist and biologist to adapt and follow in vivo these multimodal objects is required. However, the multidisciplinary approaches implemented in numerous research programs will be able to overcome these limitations.

Funding

This research was funded by CNRS, grant number PICS07354.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Bushberg, J.T.; Seibert, J.A.; Leidholdt, E.M., Jr.; Boone, J.M. The Essential Physics of Medical Imaging, 2nd ed.; Lippincott Williams & Wilkins: Philadelphia, UK, 2011. [Google Scholar]
  2. Pagel, M.D. The Hope and Hype of Multimodality Imaging Contrast Agents. Nanomedicine 2011, 6, 945–948. [Google Scholar] [CrossRef] [PubMed]
  3. Caschera, L.; Lazzara, A.; Piergallini, L.; Ricci, D.; Tuscano, B.; Vanzulli, A. Contrast Agents in Diagnostic Imaging: Present and Future. Pharm. Res. 2016, 110, 65–75. [Google Scholar] [CrossRef] [PubMed]
  4. Bigall, N.C.; Parak, W.J.; Dorfs, D. Fluorescent, Magnetic and Plasmonic—Hybrid Multifunctional Colloidal Nano Objects. Nano Today 2012, 7, 282–296. [Google Scholar] [CrossRef]
  5. Corr, S.A.; Rakovich, Y.P.; Gun’Ko, Y.K. Multifunctional Magnetic-Fluorescent Nanocomposites for Biomedical Applications. Nanoscale Res. Lett. 2008, 3, 87–104. [Google Scholar] [CrossRef] [Green Version]
  6. Shen, Z.; Wu, A.; Chen, X. Iron Oxide Nanoparticle Based Contrast Agents for Magnetic Resonance Imaging. Mol. Pharm. 2017, 14, 1352–1364. [Google Scholar] [CrossRef]
  7. Amoroso, A.J.; Pope, S.J.A. Using Lanthanide Ions in Molecular Bioimaging. Chem. Soc. Rev. 2015, 44, 4723–4742. [Google Scholar] [CrossRef] [Green Version]
  8. Ersoy, H.; Rybicki, F.J. Biochemical Safety Profiles of Gadolinium-Based Extracellular Contrast Agents and Nephrogenic Systemic Fibrosis. J. Magn. Reson. Imaging 2007, 26, 1190–1197. [Google Scholar] [CrossRef] [Green Version]
  9. Perazella, M.A. Current Status of Gadolinium Toxicity in Patients with Kidney Disease. Clin. J. Am. Soc. Nephrol. 2009, 4, 461–469. [Google Scholar] [CrossRef]
  10. Food and Drug Administration. Available online: https://www.fda.gov/downloads/Drugs/DrugSafety/UCM589442.pdf (accessed on 14 November 2019).
  11. European Medicines Agency. Available online: https://www.ema.europa.eu/en/medicines/human/referrals/gadolinium-containing-contrast-agents (accessed on 14 November 2019).
  12. Wu, L.; Mendoza-Garcia, A.; Li, Q.; Sun, S. Organic Phase Syntheses of Magnetic Nanoparticles and Their Applications. Chem. Rev. 2016, 116, 10473–11512. [Google Scholar] [CrossRef]
  13. Wang, Y.-X.J. Superparamagnetic Iron Oxide Based MRI Contrast Agents: Current Status of Clinical Application. Quant. Imaging Med. Surg. 2011, 1, 35–40. [Google Scholar] [CrossRef]
  14. Gossuin, Y.; Gillis, P.; Hocq, A.; Vuong, Q.L.; Roch, A. Magnetic Resonance Relaxation Properties of Superparamagnetic Particles. WIREs Nanomed. Nanobiotechnol. 2009, 1, 299–310. [Google Scholar] [CrossRef] [PubMed]
  15. Lee, N.; Yoo, D.; Ling, D.; Cho, M.H.; Hyeon, T.; Cheon, J. Iron Oxide Based Nanoparticles for Multimodal Imaging and Magnetoresponsive Therapy. Chem. Rev. 2015, 115, 10637–10689. [Google Scholar] [CrossRef] [PubMed]
  16. Shin, T.H.; Choi, Y.; Kim, S.; Cheon, J. Recent Advances in Magnetic Nanoparticle-Based Multi-Modal Imaging. Chem. Soc. Rev. 2015, 44, 4501–4516. [Google Scholar] [CrossRef] [PubMed]
  17. Qin, J.; Liu, Q.; Zhang, J.; Chen, J.; Chen, S.; Zhao, Y.; Du, J. Rationally Separating the Corona and Membrane Functions of Polymer Vesicles for Enhanced T2 MRI and Drug Delivery. ACS Appl. Mater. Interfaces 2015, 7, 14043–14052. [Google Scholar] [CrossRef] [PubMed]
  18. Sanson, C.; Diou, O.; Thévenot, J.; Ibarboure, E.; Soum, A.; Brûlet, A.; Miraux, S.; Thiaudière, E.; Tan, S.; Brisson, A.; et al. Doxorubicin Loaded Magnetic Polymersomes: Theranostic Nanocarriers for MR Imaging and Magneto-Chemotherapy. ACS Nano 2011, 5, 1122–1140. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  19. Martina, M.S.; Fortin, J.P.; Ménager, C.; Clément, O.; Barratt, G.; Grabielle-Madelmont, C.; Gazeau, F.; Cabuil, V.; Lesieur, S. Generation of Superparamagnetic Liposomes Revealed as Highly Efficient MRI Contrast Agents for in Vivo Imaging. J. Am. Chem. Soc. 2005, 127, 10676–10685. [Google Scholar] [CrossRef]
  20. Ren, T.; Liu, Q.; Lu, H.; Liu, H.; Zhang, X.; Du, J. Multifunctional Polymer Vesicles for Ultrasensitive Magnetic Resonance Imaging and Drug Delivery. J. Mater. Chem. 2012, 22, 12329–12338. [Google Scholar] [CrossRef]
  21. Pothayee, N.; Balasubramaniam, S.; Pothayee, N.; Jain, N.; Hu, N.; Lin, Y.; Davis, R.M.; Sriranganathan, N.; Koretsky, A.P.; Riffle, J.S. Magnetic Nanoclusters with Hydrophilic Spacing for Dual Drug Delivery and Sensitive Magnetic Resonance Imaging. J. Mater. Chem. B 2013, 1, 1142–1149. [Google Scholar] [CrossRef]
  22. Hickey, R.J.; Koski, J.; Meng, X.; Riggleman, R.A.; Zhang, P.; Park, S.-J.J. Size-Controlled Self-Assembly of Superparamagnetic Polymersomes. ACS Nano 2014, 8, 495–502. [Google Scholar] [CrossRef] [Green Version]
  23. Mikhaylov, G.; Mikac, U.; Magaeva, A.A.; Itin, V.I.; Naiden, E.P.; Psakhye, I.; Babes, L.; Reinheckel, T.; Peters, C.; Zeiser, R.; et al. Ferri-Liposomes as an MRI-Visible Drug-Delivery System for Targeting Tumours and Their Microenvironment. Nat. Nanotechnol. 2011, 6, 594–602. [Google Scholar] [CrossRef]
  24. Bleul, R.; Thiermann, R.; Marten, G.U.; House, M.J.; Pierre, T.G.S.; Häfeli, U.O.; Maskos, M. Continuously Manufactured Magnetic Polymersomes-a Versatile Tool (Not Only) for Targeted Cancer Therapy. Nanoscale 2013, 5, 11385–11393. [Google Scholar] [CrossRef] [PubMed]
  25. Prashant, C.; Dipak, M.; Yang, C.T.; Chuang, K.H.; Jun, D.; Feng, S.S. Superparamagnetic Iron Oxide—Loaded Poly (Lactic Acid)-d-α-Tocopherol Polyethylene Glycol 1000 Succinate Copolymer Nanoparticles as MRI Contrast Agent. Biomaterials 2010, 31, 5588–5597. [Google Scholar] [CrossRef] [PubMed]
  26. Ai, H.; Flask, C.; Weinberg, B.; Farrell, D.; Pagel, M.D.; Ai, H.; Shuai, X.-T.; Gao, J.; Duerk, J.; Gao, J. Magnetite-Loaded Polymeric Micelles as Ultrasensitive Magnetic-Resonance Probes. Adv. Mater. 2005, 17, 1949–1952. [Google Scholar] [CrossRef]
  27. Roch, A.; Gossuin, Y.; Muller, R.N.; Gillis, P. Superparamagnetic Colloid Suspensions: Water Magnetic Relaxation and Clustering. J. Magn. Magn. Mater. 2005, 293, 532–539. [Google Scholar] [CrossRef]
  28. Tanaka, K.; Narita, A.; Kitamura, N.; Uchiyama, W.; Morita, M.; Inubushi, T.; Chujo, Y. Preparation for Highly Sensitive MRI Contrast Agents Using Core/Shell Type Nanoparticles Consisting of Multiple SPIO Cores with Thin Silica Coating. Langmuir 2010, 26, 11759–11762. [Google Scholar] [CrossRef]
  29. Ebert, S.; Bannwarth, M.B.; Musyanovych, A.; Landfester, K.; Münnemann, K. How Morphology Influences Relaxivity—Comparative Study of Superparamagnetic Iron Oxide-Polymer Hybrid Nanostructures. Contrast Media Mol. Imaging 2015, 10, 456–464. [Google Scholar] [CrossRef]
  30. Bulte, J.W.M.; De Cuyper, M. Magnetoliposomes as Contrast Agents. Methods Enzymol. 2003, 373, 175–198. [Google Scholar] [CrossRef]
  31. Pflipsen, C.; Forge, D.; Benali, S.; Gossuin, Y. Improved Stability and Relaxivity of a Commercial Magnetic Ferrofluid. J. Phys. Chem. C 2013, 117, 20919–20926. [Google Scholar] [CrossRef]
  32. Hobson, N.J.; Weng, X.; Siow, B.; Veiga, C.; Ashford, M. Clustering Superparamagnetic Iron Oxide Nanoparticles Produces Organ-Targeted High-Contrast Magnetic Resonance Images. Nanomedicine 2019, 14, 1135–1152. [Google Scholar] [CrossRef]
  33. Yang, J.; Lee, C.H.; Ko, H.J.; Suh, J.S.; Yoon, H.G.; Lee, K.; Huh, Y.M.; Haam, S. Multifunctional Magneto-Polymeric Nanohybrids for Targeted Detection and Synergistic Therapeutic Effects on Breast Cancer. Angew. Chem. Int. Ed. 2007, 46, 8836–8839. [Google Scholar] [CrossRef]
  34. Meledandri, C.J.; Ninjbadgar, T.; Brougham, D.F. Size-Controlled Magnetoliposomes with Tunable Magnetic Resonance Relaxation Enhancements. J. Mater. Chem. 2011, 21, 214–222. [Google Scholar] [CrossRef]
  35. Taboada, E.; Solanas, R.; Rodríguez, E.; Weissleder, R.; Roig, A. Supercritical-Fluid-Assisted One-Pot Synthesis of Biocompatible Core(γ-Fe2O3)/Shell(SiO2) Nanoparticles as High Relaxivity T2-Contrast Agents for Magnetic Resonance Imaging. Adv. Funct. Mater. 2009, 19, 2319–2324. [Google Scholar] [CrossRef]
  36. Poselt, E.; Kloust, H.; Tromsdorf, U.; Janschel, M.; Hahn, C.; Masslo, C.; Weller, H. Relaxivity Optimization of a PEGylated Iron-Oxide-Based Negative Magnetic Resonance Contrast Agent for T2-Weighted Spin- Echo Imaging. ACS Nano 2012, 6, 1619–1624. [Google Scholar] [CrossRef] [PubMed]
  37. Arosio, P.; Thévenot, J.; Orlando, T.; Orsini, F.; Corti, M.; Mariani, M.; Bordonali, L.; Innocenti, C.; Sangregorio, C.; Oliveira, H.; et al. Hybrid Iron Oxide-Copolymer Micelles and Vesicles as Contrast Agents for MRI: Impact of the Nanostructure on the Relaxometric Properties. J. Mater. Chem. B 2013, 1, 5317–5328. [Google Scholar] [CrossRef] [Green Version]
  38. He, J.; Liu, X.; Niu, D.; Chen, J.; Qin, X.; Li, Y. Supramolecular-Based PEGylated Magnetic Hybrid Vesicles with Ultra-High Transverse Relaxivity. Appl. Mater. Today 2018, 11, 238–245. [Google Scholar] [CrossRef]
  39. Paquet, C.; De Haan, H.W.; Leek, D.M.; Lin, H.Y.; Xiang, B.; Tian, G.; Kell, A.; Simard, B. Clusters of Superparamagnetic Iron Oxide Nanoparticles Encapsulated in a Hydrogel: A Particle Architecture Generating a Synergistic Enhancement of the T2 Relaxation. ACS Nano 2011, 5, 3104–3112. [Google Scholar] [CrossRef] [Green Version]
  40. Yuan, Y.; He, Y.; Bo, R.; Ma, Z.; Wang, Z.; Dong, L.; Lin, T.Y.; Xue, X.; Li, Y. A Facile Approach to Fabricate Self-Assembled Magnetic Nanotheranostics for Drug Delivery and Imaging. Nanoscale 2018, 10, 21634–21639. [Google Scholar] [CrossRef]
  41. Berret, J.-F.; Schonbeck, N.; Gazeau, F.; El Kharrat, D.; Sandre, O.; Vacher, A.; Airiau, M. Controlled Clustering of Superparamagnetic Nanoparticles Using Block Copolymers: Design of New Contrast Agents for Magnetic Resonance Imaging. J. Am. Chem. Soc. 2006, 128, 1755–1761. [Google Scholar] [CrossRef]
  42. Schmidtke, C.; Eggers, R.; Zierold, R.; Feld, A.; Kloust, H.; Wolter, C.; Ostermann, J.; Merkl, J.P.; Schotten, T.; Nielsch, K.; et al. Polymer-Assisted Self-Assembly of Superparamagnetic Iron Oxide Nanoparticles into Well-Defined Clusters: Controlling the Collective Magnetic Properties. Langmuir 2014, 30, 11190–11196. [Google Scholar] [CrossRef]
  43. Liu, Q.; Song, L.; Chen, S.; Gao, J.; Zhao, P.; Du, J. A Superparamagnetic Polymersome with Extremely High T2 Relaxivity for MRI and Cancer-Targeted Drug Delivery. Biomaterials 2017, 114, 23–33. [Google Scholar] [CrossRef]
  44. Schaller, V.; Wahnström, G.; Sanz-Velasco, A.; Enoksson, P.; Johansson, C. Monte Carlo Simulation of Magnetic Multi-Core Nanoparticles. J. Magn. Magn. Mater. 2009, 321, 1400–1403. [Google Scholar] [CrossRef]
  45. Schaller, V.; Wahnström, G.; Sanz-Velasco, A.; Gustafsson, S.; Olsson, E.; Enoksson, P.; Johansson, C. Effective Magnetic Moment of Magnetic Multicore Nanoparticles. Phys. Rev. B 2009, 80, 092406. [Google Scholar] [CrossRef]
  46. Pedrosa, S.S.; Martins, S.M.S.B.; Souza, R.M.; Dantas, J.T.S.; Souza, C.M.; Rebouças, G.O.G.; de Araújo, J.M.; Dantas, A.L.; Carriço, A.S. Dipolar Effects on the Magnetic Phases of Superparamagnetic Clusters. J. Appl. Phys. 2018, 123, 233902. [Google Scholar] [CrossRef]
  47. Allia, P.; Tiberto, P.; Coisson, M.; Chiolerio, A.; Celegato, F.; Vinai, F.; Sangermano, M.; Suber, L.; Marchegiani, G. Evidence for Magnetic Interactions among Magnetite Nanoparticles Dispersed in Photoreticulated PEGDA-600 Matrix. J. Nanopart. Res. 2011, 13, 5615–5626. [Google Scholar] [CrossRef]
  48. Bae, C.J.; Angappane, S.; Park, J.G.; Lee, Y.; Lee, J.; An, K.; Hyeon, T. Experimental Studies of Strong Dipolar Interparticle Interaction in Monodisperse Fe3O4 Nanoparticles. Appl. Phys. Lett. 2007, 91, 102502. [Google Scholar] [CrossRef] [Green Version]
  49. Cha, J.; Kwon, Y.S.; Yoon, T.J.; Lee, J.K. Relaxivity Control of Magnetic Nanoclusters for Efficient Magnetic Relaxation Switching Assay. Chem. Commun. 2013, 49, 457–459. [Google Scholar] [CrossRef]
  50. Vuong, Q.L.; Berret, J.-F.; Fresnais, J.; Gossuin, Y.; Sandre, O. A Universal Scaling Law to Predict the Efficiency of Magnetic Nanoparticles as MRI T2-Contrast Agents. Adv. Healthc. Mater. 2012, 1, 502–512. [Google Scholar] [CrossRef] [Green Version]
  51. Massart, R.; Cabuil, V. Synthèse En Milieu Alcalin de Magnétite Colloïdale: Contrôle Du Rendement et de La Taille Des Particules. J. Chim. Phys. 1987, 84, 967–973. [Google Scholar] [CrossRef]
  52. Bacri, J.C.; Perzynski, R.; Salin, D.; Cabuil, V.; Massart, R. Magnetic Colloidal Properties of Ionic Ferrofluids. J. Magn. Magn. Mater. 1986, 62, 36–46. [Google Scholar] [CrossRef]
  53. Gribanov, N.M.; Bibik, E.E.; Buzunov, O.V.; Naumov, V.N. Physico-Chemical Regularities of Obtaining Highly Dispersed Magnetite by the Method of Chemical Condensation. J. Magn. Magn. Mater. 1990, 85, 7–10. [Google Scholar] [CrossRef]
  54. Lefebure, S.; Dubois, E.; Cabuil, V.; Neveu, S.; Massart, R.; Lefebure, S.; Dubois, E.; Neveu, S. Monodisperse Magnetic Nanoparticles: Preparation and Dispersion in Water and Oils. J. Mater. Res. 1998, 13, 2975–2981. [Google Scholar] [CrossRef]
  55. Lucas, I.T.; Durand-Vidal, S.; Dubois, E.; Chevalet, J.; Turq, P. Surface Charge Density of Maghemite Nanoparticles: Role of Electrostatics in the Proton Exchange. J. Phys. Chem. C 2007, 111, 18568–18576. [Google Scholar] [CrossRef] [Green Version]
  56. Bacri, J.C.; Perzynski, R.; Salin, D.; Cabuil, V.; Massart, R. Ionic Ferrofluids: A Crossing of Chemistry and Physics. J. Magn. Magn. Mater. 1990, 85, 27–32. [Google Scholar] [CrossRef]
  57. Bee, A.; Massart, R.; Neveu, S. Synthesis of Very Fine Maghemite Particles. J. Magn. Magn. Mater. 1995, 149, 6–9. [Google Scholar] [CrossRef]
  58. Chanteau, B.; Fresnais, J.; Berret, J.F. Electrosteric Enhanced Stability of Functional Sub-10 Nm Cerium and Iron Oxide Particles in Cell Culture Medium. Langmuir 2009, 25, 9064–9070. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  59. Berret, J.F.; Sandre, O.; Mauger, A. Size Distribution of Superparamagnetic Particles Determined by Magnetic Sedimentation. Langmuir 2007, 23, 2993–2999. [Google Scholar] [CrossRef] [PubMed]
  60. Gnanaprakash, G.; Philip, J.; Jayakumar, T.; Raj, B. Effect of Digestion Time and Alkali Addition Rate on Physical Properties of Magnetite Nanoparticles. J. Phys. Chem. B 2007, 111, 7978–7986. [Google Scholar] [CrossRef]
  61. Santoyo Salazar, J.; Perez, L.; De Abril, O.; Truong Phuoc, L.; Ihiawakrim, D.; Vazquez, M.; Greneche, J.M.; Begin-Colin, S.; Pourroy, G. Magnetic Iron Oxide Nanoparticles in 10–40 Nm Range: Composition in Terms of Magnetite/Maghemite Ratio and Effect on the Magnetic Properties. Chem. Mater. 2011, 23, 1379–1386. [Google Scholar] [CrossRef]
  62. Hyeon, T.; Lee, S.S.; Park, J.; Chung, Y.; Na, H.B. Synthesis of Highly Crystalline and Monodisperse Maghemite Nanocrystallites without a Size-Selection Process. J. Am. Chem. Soc. 2001, 123, 12798–12801. [Google Scholar] [CrossRef]
  63. Sun, S.; Zeng, H. Size-Controlled Synthesis of Magnetite Nanoparticles. J. Am. Chem. Soc. 2002, 124, 8204–8205. [Google Scholar] [CrossRef]
  64. Sun, S.; Zeng, H.; Robinson, D.B.; Raoux, S.; Rice, P.M.; Wang, S.X.; Li, G. Monodisperse MFe2O4 (M = Fe, Co, Mn) Nanoparticles. J. Am. Chem. Soc. 2004, 126, 273–279. [Google Scholar] [CrossRef] [PubMed]
  65. Yu, W.W.; Falkner, J.C.; Yavuz, C.T.; Colvin, V.L. Synthesis of Monodisperse Iron Oxide Nanocrystals by Thermal Decomposition of Iron Carboxylate Salts. Chem. Commun. 2004, 20, 2306–2307. [Google Scholar] [CrossRef] [PubMed]
  66. Park, J.; An, K.; Hwang, Y.; Park, J.-G.; Noh, H.-J.; Kim, J.-Y.; Park, J.-H.; Hwang, N.-M.; Hyeon, T. Ultra-Large-Scale Syntheses of Monodisperse Nanocrystals. Nat. Mater. 2004, 3, 891–895. [Google Scholar] [CrossRef] [PubMed]
  67. Park, J.; Lee, E.; Hwang, N.-M.; Kang, M.; Kim, S.C.; Hwang, Y.; Hyeon, T. One-Nanometer-Scale Size-Controlled Synthesis of Monodisperse Magnetic Iron Oxide Nanoparticles. Angew. Chem. Int. Ed. 2005, 44, 2873–2877. [Google Scholar] [CrossRef] [PubMed]
  68. Demortière, A.; Panissod, P.; Pichon, B.P.; Pourroy, G.; Guillon, D.; Donnio, B.; Bégin-Colin, S. Size-Dependent Properties of Magnetic Iron Oxide Nanocrystals. Nanoscale 2011, 3, 225–232. [Google Scholar] [CrossRef] [PubMed]
  69. Lartigue, L.; Innocenti, C.; Kalaivani, T.; Awwad, A.; Sanchez Duque, M.D.M.; Guari, Y.; Arosio, P. Water-Dispersible Sugar-Coated Iron Oxide Nanoparticles. An Evaluation of Their Relaxometric and Magnetic Hyperthermia Properties. J. Am. Chem. Soc. 2011, 133, 10459–10472. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  70. Lin, C.-R.; Chiang, R.-K.; Wang, J.-S.; Sung, T.-W. Magnetic Properties of Monodisperse Iron Oxide Nanoparticles. J. Appl. Phys. 2006, 99, 08N710. [Google Scholar] [CrossRef]
  71. Hugounenq, P.; Levy, M.; Alloyeau, D.; Lartigue, L.; Dubois, E.; Cabuil, V.; Ricolleau, C.; Roux, S.; Wilhelm, C.; Gazeau, F.; et al. Iron Oxide Monocrystalline Nanoflowers for Highly Efficient Magnetic Hyperthermia. J. Phys. Chem. C 2012, 116, 15702–15712. [Google Scholar] [CrossRef]
  72. Cheng, C.; Xu, F.; Gu, H. Facile Synthesis and Morphology Evolution of Magnetic Iron Oxide Nanoparticles in Different Polyol Processes. New J. Chem. 2011, 35, 1072. [Google Scholar] [CrossRef]
  73. Ge, J.; Hu, Y.; Biasini, M.; Beyermann, W.P.; Yin, Y. Superparamagnetic Magnetite Colloidal Nanocrystal Clusters. Angew. Chem. Int. Ed. 2007, 46, 4342–4345. [Google Scholar] [CrossRef]
  74. Lartigue, L.; Hugounenq, P.; Alloyeau, D.; Clarke, S.P.; Levy, M.; Bazzi, R.; Brougham, D.F.; Wilhelm, C.; Gazeau, F.; Lévy, M.; et al. Cooperative Organization in Iron Oxide Multi-Core Nanoparticles Potentiates Their Efficiency as Heating Mediators and MRI Contrast Agents. ACS Nano 2012, 6, 10935–10949. [Google Scholar] [CrossRef] [PubMed]
  75. Kostopoulou, A.; Velu, S.K.P.; Thangavel, K.; Orsini, F.; Brintakis, K.; Psycharakis, S.; Ranella, A.; Bordonali, L.; Lappas, A.; Lascialfari, A. Colloidal Assemblies of Oriented Maghemite Nanocrystals and Their NMR Relaxometric Properties. Dalt. Trans. 2014, 43, 8395–8404. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Ettinger, A.; Wittmann, T. Fluorescence Live Cell Imaging. Methods Cell Biol. 2014, 123, 77–94. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  77. Sanderson, M.J.; Smith, I.; Parker, I.; Bootman, M.D. Fluorescence Microscopy. Cold Spring Harb. Protoc. 2014, 1042–1065. [Google Scholar] [CrossRef] [Green Version]
  78. Hell, S.W. Toward Fluorescence Nanoscopy. Nat. Biotechnol. 2003, 21, 1347–1355. [Google Scholar] [CrossRef]
  79. Palmer, A.E.; Tsien, R.Y. Measuring Calcium Signaling Using Genetically Targetable Fluorescent Indicators. Nat. Protoc. 2006, 1, 1057–1065. [Google Scholar] [CrossRef]
  80. Paige, J.S.; Nguyen-Duc, T.; Song, W.; Jaffrey, S.R. Fluorescence Imaging of Cellular Metabolites with RNA. Science 2012, 335, 1194. [Google Scholar] [CrossRef] [Green Version]
  81. Weiss, S. Fluorescence Spectroscopy of Single Biomolecules. Science 1999, 283, 1676–1683. [Google Scholar] [CrossRef] [Green Version]
  82. Anderson, N.G.; Mann, M.; Meng, C.K.; Wong, S.F.; Hillenkamp, F.; Goodlet, D.R.; Mann, R.M.; Glish, G.L.; Mcluckey, S.A.; Kaiser, R.E. The Fluorescent Toolbox for Assessing Protein Location and Function. Science 2006, 312, 217–224. [Google Scholar] [CrossRef] [Green Version]
  83. Resch-Genger, U.; Grabolle, M.; Cavaliere-Jaricot, S.; Nitschke, R.; Nann, T. Quantum Dots versus Organic Dyes as Fluorescent Labels. Nat. Methods 2008, 5, 763–775. [Google Scholar] [CrossRef]
  84. Williams, A.T.R.; Winfield, S.A.; Miller, J.N. Relative Fluorescence Quantum Yields Using a Computer-Controlled Luminescence Spectrometer. Analyst 1983, 108, 1067–1071. [Google Scholar] [CrossRef]
  85. Wu, C.; Szymanski, C.; Cain, Z.; McNeill, J. Conjugated Polymer Dots for Multiphoton Fluorescence Imaging. J. Am. Chem. Soc. 2007, 129, 12904–12905. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Smith, A.M.; Duan, H.; Mohs, A.M.; Nie, S. Bioconjugated Quantum Dots for in Vivo Molecular and Cellular Imaging. Adv. Drug Deliv. Rev. 2008, 60, 1226–1240. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  87. Sharma, P.; Brown, S.; Walter, G.; Santra, S.; Moudgil, B. Nanoparticles for Bioimaging. Adv. Colloid Interface Sci. 2006, 123–126, 471–485. [Google Scholar] [CrossRef] [PubMed]
  88. Terai, T.; Nagano, T. Small-Molecule Fluorophores and Fluorescent Probes for Bioimaging. Pflugers Arch. Eur. J. Physiol. 2013, 465, 347–359. [Google Scholar] [CrossRef]
  89. Wang, L.; Frei, M.S.; Salim, A.; Johnsson, K. Small-Molecule Fluorescent Probes for Live-Cell Super-Resolution Microscopy. J. Am. Chem. Soc. 2019, 141, 2770–2781. [Google Scholar] [CrossRef]
  90. Johnson, I.; Spence, M.T.Z. The Molecular Probe® Handbook: A Guide to Fluorescent Probes and Labeling Technologies, 11th ed.; Life Technologies Corporation: Carlsbad, CA, USA, 2010. [Google Scholar]
  91. Johnson, L.V.; Walsh, M.L.; Chen, L.B. Localization of Mitochondria in Living Cells with Rhodamine 123. Proc. Natl. Acad. Sci. USA 1980, 77, 990–994. [Google Scholar] [CrossRef] [Green Version]
  92. Anderson, R.G. A View of Acidic Intracellular Compartments. J. Cell Biol. 2004, 106, 539–543. [Google Scholar] [CrossRef] [Green Version]
  93. Crissman, H.A.; Hirons, G.T. Staining of DNA in Live and Fixed Cells. Methods Cell Biol. 1994, 41, 195–209. [Google Scholar] [CrossRef]
  94. Ishow, E.; Brosseau, A.; Clavier, G.; Nakatani, K.; Tauc, P.; Neveu, S.; Sandre, O.; Léaustic, A. Multicolor Emission of Small Molecule-Based Amorphous Thin Films and Nanoparticles with a Single Excitation Wavelength. Chem. Mater. 2008, 20, 6597–6599. [Google Scholar] [CrossRef] [Green Version]
  95. Patra, A.; Chandaluri, C.G.; Radhakrishnan, T.P. Optical Materials Based on Molecular Nanoparticles. Nanoscale 2012, 4, 343–359. [Google Scholar] [CrossRef] [PubMed]
  96. Lei, T.; Pei, J. Solution-Processed Organic Nano- and Micro-Materials: Design Strategy, Growth Mechanism and Applications. J. Mater. Chem. 2012, 22, 785–798. [Google Scholar] [CrossRef]
  97. Cui, Q.H.; Zhao, Y.S.; Yao, J. Controlled Synthesis of Organic Nanophotonic Materials with Specific Structures and Compositions. Adv. Mater. 2014, 26, 6852–6870. [Google Scholar] [CrossRef] [PubMed]
  98. Zhao, Y.S.; Fu, H.; Peng, A.; Ma, Y.; Xiao, D.; Yao, J. Low-Dimensional Nanomaterials Based on Small Organic Molecules: Preparation and Optoelectronic Properties. Adv. Mater. 2008, 20, 2859–2876. [Google Scholar] [CrossRef]
  99. Fischer, I.; Kaeser, A.; Peters-Gumbs, M.A.M.; Schenning, A.P.H.J. Fluorescent π-Conjugated Polymer Dots versus Self-Assembled Small-Molecule Nanoparticles: What’s the Difference? Chem. A Eur. J. 2013, 19, 10928–10934. [Google Scholar] [CrossRef]
  100. Gaiduk, A.; Yorulmaz, M.; Ishow, E.; Orrit, M. Absorption, Luminescence, and Sizing of Organic Dye Nanoparticles and of Patterns Formed Upon Dewetting. ChemPhysChem 2012, 13, 946–951. [Google Scholar] [CrossRef]
  101. Faucon, A.; Benhelli-Mokrani, H.; Córdova, L.A.W.; Brulin, B.; Heymann, D.; Hulin, P.; Nedellec, S.; Ishow, E. Are Fluorescent Organic Nanoparticles Relevant Tools for Tracking Cancer Cells or Macrophages? Adv. Healthc. Mater. 2015, 4, 2727–2734. [Google Scholar] [CrossRef]
  102. Day, R.N.; Davidson, M.W. The Fluorescent Protein Palette: Tools for Cellular Imaging. Chem. Soc. Rev. 2009, 38, 2887–2921. [Google Scholar] [CrossRef] [Green Version]
  103. Shimomura, O. The discovery of aequorin and green fluorescent protein. J. Microsc. 2005, 217, 3–15. [Google Scholar] [CrossRef]
  104. Chalfie, M.; Tu, Y.; Euskirchen, G.; Ward William, W.; Prasher Douglas, C. Green Fluorescent Protein as a Marker for Gene Expression. Science 1994, 263, 802–805. [Google Scholar] [CrossRef] [Green Version]
  105. Shimomura, O. Structure of the Chromophore of Aequorea Green Fluorescent Protein. FEBS Lett. 1979, 104, 220–222. [Google Scholar] [CrossRef] [Green Version]
  106. Yang, F.; Moss, L.G.; Phillips, G.N., Jr. The Molecular Structure of Green Fluorescent Protein. Nat. Biotechnol. 1996, 14, 1246–1251. [Google Scholar] [CrossRef] [PubMed]
  107. Tsien, R.Y. The Green Fluorescent Protein. Ann. Rev. Biochem. 1998, 67, 509–544. [Google Scholar] [CrossRef] [PubMed]
  108. Jensen, E.C. Use of Fluorescent Probes: Their Effect on Cell Biology and Limitations. Anat. Rec. (Hoboken) 2012, 295, 2031–2036. [Google Scholar] [CrossRef] [PubMed]
  109. Chan, Y.H.; Wu, C.; Ye, F.; Jin, Y.; Smith, P.B.; Chiu, D.T. Development of Ultrabright Semiconducting Polymer Dots for Ratiometric PH Sensing. Anal. Chem. 2011, 83, 1448–1455. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  110. Tuncel, D.; Demir, H.V. Conjugated Polymer Nanoparticles. Nanoscale 2010, 2, 484–494. [Google Scholar] [CrossRef]
  111. Burnham, D.R.; Zeigler, M.; Wu, C.; McNeill, J.D.; Yu, J.; Chiu, D.T.; Schneider, T.; Schiro, P.G. Bioconjugation of Ultrabright Semiconducting Polymer Dots for Specific Cellular Targeting. J. Am. Chem. Soc. 2010, 132, 15410–15417. [Google Scholar] [CrossRef] [Green Version]
  112. Wu, C.; Bull, B.; Szymanski, C.; Christensen, K.; McNeill, J. Multicolor Conjugated Polymer Dots for Biological Fluorescence Imaging. ACS Nano 2008, 2, 2415–2423. [Google Scholar] [CrossRef]
  113. Wu, C.; Chiu, D.T. Highly Fluorescent Semiconducting Polymer Dots for Biology and Medicine. Angew. Chem. Int. Ed. 2013, 52, 3086–3109. [Google Scholar] [CrossRef] [Green Version]
  114. Feng, L.; Zhu, C.; Yuan, H.; Liu, L.; Lv, F.; Wang, S. Conjugated Polymer Nanoparticles: Preparation, Properties, Functionalization and Biological Applications. Chem. Soc. Rev. 2013, 42, 6620–6633. [Google Scholar] [CrossRef]
  115. Zhu, C.; Liu, L.; Yang, Q.; Lv, F.; Wang, S. Water-Soluble Conjugated Polymers for Imaging, Diagnosis, and Therapy. Chem. Rev. 2012, 112, 4687–4735. [Google Scholar] [CrossRef] [PubMed]
  116. Wu, C.; Hansen, S.J.; Hou, Q.; Yu, J.; Zeigler, M.; Jin, Y.; Burnham, D.R.; McNeill, J.D.; Olson, J.M.; Chiu, D.T. Design of Highly Emissive Polymer Dot Bioconjugates for in Vivo Tumor Targeting. Angew. Chem. Int. Ed. 2011, 50, 3430–3434. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  117. Chen, X.; Liu, Z.; Li, R.; Shan, C.; Zeng, Z.; Xue, B.; Yuan, W.; Mo, C.; Xi, P.; Wu, C.; et al. Multicolor Super-Resolution Fluorescence Microscopy with Blue and Carmine Small Photoblinking Polymer Dots. ACS Nano 2017, 11, 8084–8091. [Google Scholar] [CrossRef] [PubMed]
  118. Dubertret, B.; Skourides, P.; Norris, D.J.; Noireaux, V.; Brivanlou, A.H.; Libchaber, A. In Vivo Imaging of Quantum Dots Encapsulated in Phospholipid Micelles. Science 2002, 298, 1759–1762. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  119. Larson, D.R.; Zipfel, W.R.; Williams, R.M.; Clark, S.W.; Bruchez, M.P.; Wise, F.W.; Webb, W.W. Water-Soluble Quantum Dots for Multiphoton Fluorescence Imaging in Vivo. Science 2003, 300, 1434–1436. [Google Scholar] [CrossRef]
  120. Åkerman, M.E.; Chan, W.C.W.; Laakkonen, P.; Bhatia, S.N.; Ruoslahti, E. Nanocrystal Targeting in Vivo. Proc. Natl. Acad. Sci. USA 2002, 99, 12617–12621. [Google Scholar] [CrossRef] [Green Version]
  121. Bruchez, M., Jr.; Moronne, M.; Gin, P.; Weiss, S.; Alivisatos, A.P. Semiconductor Nanocrystals as Fluorescent Biological Labels. Science 1998, 281, 2013–2015. [Google Scholar] [CrossRef] [Green Version]
  122. Christodoulou, S.; Vaccaro, G.; Pinchetti, V.; De Donato, F.; Grim, J.Q.; Casu, A.; Genovese, A.; Vicidomini, G.; Diaspro, A.; Brovelli, S.; et al. Synthesis of Highly Luminescent Wurtzite CdSe/CdS Giant-Shell Nanocrystals Using a Fast Continuous Injection Route. J. Mater. Chem. C 2014, 2, 3439–3447. [Google Scholar] [CrossRef] [Green Version]
  123. Chen, O.; Zhao, J.; Chauhan, V.P.; Cui, J.; Wong, C.; Harris, D.K.; Wei, H.; Han, H.; Fukumura, D.; Jain, R.K.; et al. Compact High-Quality CdSe–CdS Core–shell Nanocrystals with Narrowemission Linewidths and Suppressed Blinking. Nat. Mater. 2013, 12, 445–451. [Google Scholar] [CrossRef] [Green Version]
  124. Zhou, J.; Yang, Y.; Zhang, C.Y. Toward Biocompatible Semiconductor Quantum Dots: From Biosynthesis and Bioconjugation to Biomedical Application. Chem. Rev. 2015, 115, 11669–11717. [Google Scholar] [CrossRef]
  125. Hoshino, A.; Fujioka, K.; Oku, T.; Suga, M. Physicochemical Properties and Cellular Toxicity of Nanocrystal Quantum Dots Depend on Their Surface Modification. Nano Lett. 2004, 4, 2163–2169. [Google Scholar] [CrossRef]
  126. Derfus, A.M.; Chan, W.C.W.; Bhatia, S.N. Probing the Cytotoxicity of Semiconductor Quantum Dots. Nano Lett. 2004, 4, 2163–2169. [Google Scholar] [CrossRef] [Green Version]
  127. Hemmer, E.; Benayas, A.; Légaré, F.; Vetrone, F. Exploiting the Biological Windows: Current Perspectives on Fluorescent Bioprobes Emitting above 1000 nm. Nanoscale Horiz. 2016, 1, 168–184. [Google Scholar] [CrossRef]
  128. Xi, L.; Jiang, H. Image-Guided Surgery Using Multimodality Strategy and Molecular Probes. WIREs Nanomed. Nanobiotechnol. 2016, 8, 46–60. [Google Scholar] [CrossRef]
  129. Senders, J.T.; Muskens, I.S.; Schnoor, R.; Karhade, A.V.; Cote, D.J.; Smith, T.R.; Broekman, M.L.D. Agents for Fluorescence-Guided Glioma Surgery: A Systematic Review of Preclinical and Clinical Results. Acta Neurochir. 2016, 1, 151–167. [Google Scholar] [CrossRef] [Green Version]
  130. Bertorelle, F.; Wilhelm, C.; Roger, J.; Gazeau, F.; Ménager, C.; Cabuil, V. Fluorescence-Modified Superparamagnetic Nanoparticles: Intracellular Uptake and Use in Cellular Imaging. Langmuir 2006, 22, 5385–5391. [Google Scholar] [CrossRef]
  131. Alcantara, D.; Guo, Y.; Yuan, H.; Goergen, C.J.; Chen, H.H.; Cho, H.; Sosnovik, D.E.; Josephson, L. Fluorochrome-Functionalized Magnetic Nanoparticles for High-Sensitivity Monitoring of the Polymerase Chain Reaction by Magnetic Resonance. Angew. Chem. Int. Ed. 2012, 51, 6904–6907. [Google Scholar] [CrossRef]
  132. Yen, S.K.; Jańczewski, D.; Lakshmi, J.L.; Dolmanan, S.B.; Tripathy, S.; Ho, V.H.B.; Vijayaragavan, V.; Hariharan, A.; Padmanabhan, P.; Bhakoo, K.K.; et al. Design and Synthesis of Polymer-Functionalized NIR Fluorescent Dyes-Magnetic Nanoparticles for Bioimaging. ACS Nano 2013, 7, 6796–6805. [Google Scholar] [CrossRef]
  133. Santra, S.; Kaittanis, C.; Grimm, J.; Perez, J.M. Drug/Dye-Loaded, Multifunctional Iron Oxide Nanoparticles for Combined Targeted Cancer Therapy and Dual Optical/Magnetic Resonance Imaging. Small 2009, 5, 1862–1868. [Google Scholar] [CrossRef]
  134. Redl, F.X.; Cho, K.S.; Murray, C.B.; O’Brien, S. Three-Dimensional Binary Superlattices of Magnetic Nanocrystals and Semiconductor Quantum Dots. Nature 2003, 423, 968–971. [Google Scholar] [CrossRef]
  135. Cho, M.; Contreras, E.Q.; Lee, S.S.; Jones, C.J.; Jang, W.; Colvin, V.L. Characterization and Optimization of the Fluorescence of Nanoscale Iron Oxide/Quantum Dot Complexes. J. Phys. Chem. C 2014, 118, 14606–14616. [Google Scholar] [CrossRef]
  136. Shibu, E.S.; Ono, K.; Sugino, S.; Nishioka, A.; Yasuda, A.; Shigeri, Y.; Wakida, S.I.; Sawada, M.; Biju, V. Photouncaging Nanoparticles for MRI and Fluorescence Imaging in Vitro and in Vivo. ACS Nano 2013, 7, 9851–9859. [Google Scholar] [CrossRef]
  137. Gao, J.; Zhang, W.; Huang, P.; Zhang, B.; Zhang, X.; Xu, B. Intracellular Spatial Control of Fluorescent Magnetic Nanoparticles. J. Am. Chem. Soc. 2008, 130, 3710–3711. [Google Scholar] [CrossRef]
  138. Selvan, S.T.; Patra, P.K.; Ang, C.Y.; Ying, J.Y. Synthesis of Silica-Coated Semiconductor and Magnetic Quantum Dots and Their Use in the Imaging of Live Cells. Angew. Chem. Int. Ed. 2007, 46, 2448–2452. [Google Scholar] [CrossRef]
  139. Pahari, S.K.; Olszakier, S.; Kahn, I.; Amirav, L. Magneto-Fluorescent Yolk-Shell Nanoparticles. Chem. Mater. 2018, 30, 775–780. [Google Scholar] [CrossRef]
  140. Wang, D.; He, J.; Rosenzweig, N.; Rosenzweig, Z. Superparamagnetic Fe2O3 Beads−CdSe/ZnS Quantum Dots Core−Shell Nanocomposite Particles for Cell Separation. Nano Lett. 2004, 4, 409–413. [Google Scholar] [CrossRef]
  141. Lee, J.; Lee, N.; Kim, H.; Kim, J. Mesoporous Dye-Doped Silica Nanoparticles Decorated With Multiple Magnetite Nanocrystals for Simultaneous Enhanced Magnetic Resonance Imaging, Fluorescence. J. Am. Chem. Soc. 2010, 132, 552–557. [Google Scholar] [CrossRef]
  142. Lee, J.-H.; Jun, Y.W.; Yeon, S.-I.; Shin, J.-S.; Cheon, J. Dual-Mode Nanoparticle Probes for High-Performance Magnetic Resonance and Fluorescence Imaging of Neuroblastoma. Angew. Chem. Int. Ed. 2006, 45, 8160–8162. [Google Scholar] [CrossRef]
  143. Chekina, N.; Horák, D.; Jendelová, P.; Trchová, M.; Bene, M.J.; Hrubý, M.; Herynek, V.; Turnovcová, K.; Syková, E. Fluorescent Magnetic Nanoparticles for Biomedical Applications. J. Mater. Chem. 2011, 21, 7630–7639. [Google Scholar] [CrossRef]
  144. Wang, F.; Chen, X.; Zhao, Z.; Tang, S.; Huang, X.; Lin, C.; Cai, C.; Zheng, N. Synthesis of Magnetic, Fluorescent and Mesoporous Core-Shell-Structured Nanoparticles for Imaging, Targeting and Photodynamic Therapy. J. Mater. Chem. 2011, 21, 11244–11252. [Google Scholar] [CrossRef] [Green Version]
  145. Badruddoza, A.Z.M.; Rahman, M.T.; Ghosh, S.; Hossain, M.Z.; Shi, J.; Hidajat, K.; Uddin, M.S. β-Cyclodextrin Conjugated Magnetic, Fluorescent Silica Core-Shell Nanoparticles for Biomedical Applications. Carbohydr. Polym. 2013, 95, 449–457. [Google Scholar] [CrossRef]
  146. Li, L.; Choo, E.S.G.; Liu, Z.; Ding, J.; Xue, J. Double-Layer Silica Core-Shell Nanospheres with Superparamagnetic and Fluorescent Functionalities. Chem. Phys. Lett. 2008, 461, 114–117. [Google Scholar] [CrossRef]
  147. Insin, N.; Tracy, J.B.J.; Lee, H.; Zimmer, J.P.J.; Westervelt, R.M.; Bawendi, M.G. Incorporation of Iron Oxide Nanoparticles and Quantum Dots into Silica Microspheres. ACS Nano 2008, 2, 197–202. [Google Scholar] [CrossRef] [Green Version]
  148. Sathe, T.R.; Agrawal, A.; Nie, S. Mesoporous Silica Beads Embedded with Semiconductor Quantum Dots and Iron Oxide Nanocrystals: Dual-Function Microcarriers for Optical Encoding and Magnetic Separation. Anal. Chem. 2006, 78, 5627–5632. [Google Scholar] [CrossRef]
  149. Yi, D.K.; Selvan, S.T.; Lee, S.S.; Papaefthymiou, G.C.; Kundaliya, D.; Ying, J.Y. Silica-Coated Nanocomposites of Magnetic Nanoparticles and Quantum Dots. J. Am. Chem. Soc. 2005, 127, 4990–4991. [Google Scholar] [CrossRef]
  150. Kim, J.J.; Lee, J.E.; Yu, J.H.; Kim, B.C.; An, K.; Hwang, Y.; Shin, C.H.; Park, J.G. Magnetic Fluorescent Delivery Vehicle Using Uniform Mesoporous Silica Spheres Embedded with Monodisperse Magnetic and Semiconductor Nanocrystals. J. Am. Chem. Soc. 2006, 128, 688–689. [Google Scholar] [CrossRef]
  151. He, X.; Shen, X.; Li, D.; Liu, Y.; Jia, K.; Liu, X. Dual-Mode Fluorescence and Magnetic Resonance Imaging Nanoprobe Based on Aromatic Amphiphilic Copolymer Encapsulated CdSe@CdS and Fe3O4. ACS Appl. Bio Mater. 2018, 1, 520–528. [Google Scholar] [CrossRef]
  152. Chen, O.; Riedemann, L.; Etoc, F.; Herrmann, H.; Coppey, M.; Barch, M.; Farrar, C.T.; Zhao, J.; Bruns, O.T.; Wei, H.; et al. Magneto-Fluorescent Core-Shell Supernanoparticles. Nat. Commun. 2014, 5, 1–8. [Google Scholar] [CrossRef] [Green Version]
  153. Pinkerton, N.M.; Gindy, M.E.; Calero-Ddelc, V.L.; Wolfson, T.; Pagels, R.F.; Adler, D.; Gao, D.; Li, S.; Wang, R.; Zevon, M.; et al. Single-Step Assembly of Multimodal Imaging Nanocarriers: MRI and Long-Wavelength Fluorescence Imaging. Adv. Healthc. Mater. 2015, 4, 1376–1385. [Google Scholar] [CrossRef]
  154. Das, M.; Solanki, A.; Joshi, A.; Devkar, R.; Seshadri, S.; Thakore, S. Β-Cyclodextrin Based Dual-Responsive Multifunctional Nanotheranostics for Cancer Cell Targeting and Dual Drug Delivery. Carbohydr. Polym. 2019, 206, 694–705. [Google Scholar] [CrossRef]
  155. Bixner, O.; Gal, N.; Zaba, C.; Scheberl, A.; Reimhult, E. Fluorescent Magnetopolymersomes: A Theranostic Platform to Track Intracellular Delivery. Materials 2017, 10, 1303. [Google Scholar] [CrossRef] [Green Version]
  156. Di Corato, R.; Bigall, N.C.; Ragusa, A.; Dorfs, D.; Genovese, A.; Marotta, R.; Manna, L.; Pellegrino, T. Multifunctional Nanobeads Based on Quantum Dots and Magnetic Nanoparticles: Synthesis and Cancer Cell Targeting and Sorting. ACS Nano 2011, 5, 1109–1121. [Google Scholar] [CrossRef]
  157. Ling, D.; Park, W.; Park, S.J.; Lu, Y.; Kim, K.S.; Hackett, M.J.; Kim, B.H.; Yim, H.; Jeon, Y.S.; Na, K.; et al. Multifunctional Tumor PH-Sensitive Self-Assembled Nanoparticles for Bimodal Imaging and Treatment of Resistant Heterogeneous Tumors. J. Am. Chem. Soc. 2014, 136, 5647–5655. [Google Scholar] [CrossRef]
  158. Demillo, V.G.; Zhu, X. Zwitterionic Amphiphile Coated Magnetofluorescent Nanoparticles - Synthesis, Characterization and Tumor Cell Targeting. J. Mater. Chem. B 2015, 3, 8328–8336. [Google Scholar] [CrossRef] [Green Version]
  159. Feld, A.; Merkl, J.P.; Kloust, H.; Flessau, S.; Schmidtke, C.; Wolter, C.; Ostermann, J.; Kampferbeck, M.; Eggers, R.; Mews, A.; et al. A Universal Approach to Ultrasmall Magneto-Fluorescent Nanohybrids. Angew. Chem. Int. Ed. 2015, 54, 12468–12471. [Google Scholar] [CrossRef]
  160. Béalle, G.; Di Corato, R.; Kolosnjaj-Tabi, J.; Dupuis, V.; Clément, O.; Gazeau, F.; Wilhelm, C.; Ménager, C. Ultra Magnetic Liposomes for MR Imaging, Targeting, and Hyperthermia. Langmuir 2012, 28, 11834–11842. [Google Scholar] [CrossRef]
  161. Beaune, G.; Dubertret, B.; Clément, O.; Vayssettes, C.; Cabuil, V.; Ménager, C. Giant Vesicles Containing Magnetic Nanoparticles and Quantum Dots: Feasibility and Tracking by Fiber Confocal Fluorescence Microscopy. Angew. Chem. Int. Ed. 2007, 46, 5421–5424. [Google Scholar] [CrossRef]
  162. Scheffold, A.; Miltenyi, S.; Radbruch, A. Magnetofluorescent Liposomes for Increased Sensitivity of Immunofluorescence. Immunotechnology 1995, 1, 127–137. [Google Scholar] [CrossRef]
  163. Beaune, G.; Ménager, C.; Cabuil, V. Location of Magnetic and Fluorescent Nanoparticles Encapsulated inside Giant Liposomes. J. Phys. Chem. B 2008, 112, 7424–7429. [Google Scholar] [CrossRef]
  164. Wang, G.; Zhang, X.; Liu, Y.; Hu, Z.; Mei, X.; Uvdal, K. Magneto-Fluorescent Nanoparticles with High-Intensity NIR Emission, T1- and T2-Weighted MR for Multimodal Specific Tumor Imaging. J. Mater. Chem. B 2015, 3, 3072–3080. [Google Scholar] [CrossRef]
  165. Li, K.; Ding, D.; Huo, D.; Pu, K.Y.; Thao, N.N.P.; Hu, Y.; Li, Z.; Liu, B. Conjugated Polymer Based Nanoparticles as Dual-Modal Probes for Targeted in Vivo Fluorescence and Magnetic Resonance Imaging. Adv. Funct. Mater. 2012, 22, 3107–3115. [Google Scholar] [CrossRef]
  166. Vijayan, V.M.; Ereath Beeran, A.; Shenoy, S.J.; Muthu, J.; Thomas, V. New Magneto-Fluorescent Hybrid Polymer Nanogel for Theranostic Applications. ACS Appl. Bio Mater. 2019, 2, 757–768. [Google Scholar] [CrossRef]
  167. Howes, P.; Green, M.; Bowers, A.; Parker, D.; Varma, G.; Kallumadil, M.; Hughes, M.; Warley, A.; Brain, A.; Botnar, R. Magnetic Conjugated Polymer Nanoparticles as Bimodal Imaging Agents. J. Am. Chem. Soc. 2010, 132, 9833–9842. [Google Scholar] [CrossRef] [PubMed]
  168. Faucon, A.; Maldiney, T.; Clément, O.; Hulin, P.; Nedellec, S.; Robard, M.; Gautier, N.; De Meulenaere, E.; Clays, K.; Orlando, T.; et al. Highly Cohesive Dual Nanoassemblies for Complementary Multiscale Bioimaging. J. Mater. Chem. B 2014, 2, 7747–7755. [Google Scholar] [CrossRef] [Green Version]
  169. Faucon, A.; Fresnais, J.; Brosseau, A.; Hulin, P.; Nedellec, S.; Hémez, J.; Ishow, E. Photoactive Chelating Organic Nanospheres as Central Platforms of Bimodal Hybrid Nanoparticles. J. Mater. Chem. C 2013, 1, 3879–3886. [Google Scholar] [CrossRef]
  170. Faucon, A.; Benhelli-Mokrani, H.; Fleury, F.; Dubreil, L.; Hulin, P.; Nedellec, S.; Doussineau, T.; Antoine, R.; Orlando, T.; Lascialfari, A.; et al. Tuning the Architectural Integrity of High-Performance Magneto-Fluorescent Core-Shell Nanoassemblies in Cancer Cells. J. Colloid Interface Sci. 2016, 479, 139–149. [Google Scholar] [CrossRef]
  171. Fresnais, J.; Ishow, E.; Sandre, O.; Berret, J.-F. Electrostatic Co-Assembly of Magnetic Nanoparticles and Fluorescent Nanospheres: A Versatile Approach towards Bimodal Nanorods. Small 2009, 5, 2533–2536. [Google Scholar] [CrossRef] [Green Version]
  172. Linot, C.; Poly, J.; Boucard, J.; Pouliquen, D.; Nedellec, S.; Hulin, P.; Lecouvey, M.; Marec, N.; Arosio, P.; Lascialfari, A.; et al. PEGylated Anionic Magneto Fl Uorescent Nanoassemblies: Impact of Their Interface Structure on Magnetic Resonance Imaging Contrast and Cellular Uptake. ACS Appl. Mater. Interfaces 2017, 9, 14242–14257. [Google Scholar] [CrossRef]
Nanomaterials 10 00028 g001 550
Figure 1. Fluorescence imaging vs. magnetic resonance imaging (MRI). Comparison between both imaging techniques showing their complementarity.
Figure 1. Fluorescence imaging vs. magnetic resonance imaging (MRI). Comparison between both imaging techniques showing their complementarity.
Nanomaterials 10 00028 g001
Nanomaterials 10 00028 g002 550
Figure 2. Schematic representation of the MRI. (A) Relaxation of protons under the action of an external magnetic field, B0, and radiofrequency pulse B1. (B) Return to the equilibrium position of total magnetization, M0, along the (Oz) axis and the (Oxy) plane.
Figure 2. Schematic representation of the MRI. (A) Relaxation of protons under the action of an external magnetic field, B0, and radiofrequency pulse B1. (B) Return to the equilibrium position of total magnetization, M0, along the (Oz) axis and the (Oxy) plane.
Nanomaterials 10 00028 g002
Nanomaterials 10 00028 g003 550
Figure 3. Interaction of magnetic system with water protons during an MRI experiment. (A) Schematization of the effects of magnetic nanoparticles (MNPs) on longitudinal and transverse relaxivity. (B) Probed volume by a water molecule during an MRI experiment depending on the size of the assembly. Consequence on the transverse relaxivity value and the associated model.
Figure 3. Interaction of magnetic system with water protons during an MRI experiment. (A) Schematization of the effects of magnetic nanoparticles (MNPs) on longitudinal and transverse relaxivity. (B) Probed volume by a water molecule during an MRI experiment depending on the size of the assembly. Consequence on the transverse relaxivity value and the associated model.
Nanomaterials 10 00028 g003
Nanomaterials 10 00028 g004 550
Figure 4. Self-assemblies of MNPs to obtain multi core magnetic nanoparticles. Three main organizations are noted: the encapsulation of MNPs in the core (type I), their dispersion on the shell of the nanoscale architecture (type II), or the merger of magnetic grains (type III).
Figure 4. Self-assemblies of MNPs to obtain multi core magnetic nanoparticles. Three main organizations are noted: the encapsulation of MNPs in the core (type I), their dispersion on the shell of the nanoscale architecture (type II), or the merger of magnetic grains (type III).
Nanomaterials 10 00028 g004
Nanomaterials 10 00028 g005 550
Figure 5. Description of some luminophores. (A) Chemical formula of the main families of fluorophore. (B) Structure of Pdots based on polyfluorene. (C) Cartoon illustrating the effect of QDots size on their luminescent properties.
Figure 5. Description of some luminophores. (A) Chemical formula of the main families of fluorophore. (B) Structure of Pdots based on polyfluorene. (C) Cartoon illustrating the effect of QDots size on their luminescent properties.
Nanomaterials 10 00028 g005
Nanomaterials 10 00028 g006 550
Figure 6. Schematic representation of magneto-fluorescent nanosystems.
Figure 6. Schematic representation of magneto-fluorescent nanosystems.
Nanomaterials 10 00028 g006
Table
Table 1. T2 contrast agents that have been proposed to the European and American markets. dH is the hydrodynamic diameter of the nanoparticle assembly. r2 is given at 1.5 T and 37 °C [13,14].
Table 1. T2 contrast agents that have been proposed to the European and American markets. dH is the hydrodynamic diameter of the nanoparticle assembly. r2 is given at 1.5 T and 37 °C [13,14].
NameClassedH in nm/Coatingr2 in s−1 mM−1/(r2/r1)Approval (withdrawn)Company
Endorem® or Feridex I.Vferumoxides120–180/dextran 10 kDa158 (16)1994 (2012) or 1996 (2008)Guerbet S.A. or Berlex Laboratories
Sinerem® or Combidex ®ferumoxtran-1020–40/dextran 10 kDa88 (5)n.a. (2007) or 2005 (2007)Guerbet S.A. or AMAG pharmaceuticals, Inc.
Resovist®ferucarbotran45–60/carboxydextran 1.8 kDa189 (19)2001 (2009)Bayer Healthcare
Feraheme®ferumoxytol30/semi-synthetic carbohydrate89 (6)2009AMAG pharmaceuticals, Inc.
Lumirem® or GastroMARK®ferumoxsil400/poly [N-(2-aminoethyl)-3-aminopropyl]siloxane47 (23)1993 (2014) or 1996 (2010)Guerbet S.A. or AMAG pharmaceuticals, Inc.
Table
Table 2. Properties of self-assembled magnetic nanoparticles designed as MRI contrast agent.
Table 2. Properties of self-assembled magnetic nanoparticles designed as MRI contrast agent.
TypeDispersantdcore in nmSynthesis Route (Provider)dH in nmwt% IOField/Tr2 in mM−1 s−1 (r2/r1)Ref.
IPAA-b-PS5.6TD513251.41295(n.a)[39]
6.4400378 (n.a)
10.8300561 (n.a)
15.5241555 (n.a)
IPTMC-b-PGA6.3CP50204.781 (29)[18]
4535134 (37)
4750173 (48)
5270182 (52)
IPTMC-b-PGA6–7CP12551.4171 (14)[30]
6–710951.6114 (25)
8–106733.8128 (22)
8–107950.5167 (25)
10–15875.1219 (71)
10–1514820280 (103)
IPR-PAA in organosilica matrice6TD76103642 (n.a)[31]
IPEG-b-PCL-b-PAA1.9CP14031.41108 (n.a)[17]
IPluronic® L-12110n.a. (Webcraft GmbH)1267.11.41682 (68)[41]
IDOPG or DOPC13.8CP90n.a1166 (~20)[26]
110919 (~20)
Ifolic acid-PGA-b-PCL7CP174n.a1.41612 (20)[36]
IPEG-b-poly(tert-butyl acrylate-stat-PAA6CP1754.83211 (n.a)[37]
IIaEPC and DSPE-PEG-methoxy 20007.7CP161000.47108 (3)[29]
200351116 (6)
195631130 (17)
IIbSDS9.1TD (Ferrotec)53753295 (n.a)[32]
80350 (n.a)
99410 (n.a)
IIbPCL-b-PEG4TD1712.41.525 (19)[43]
47519.5169 (58)
89738.1318 (199)
1611054.2471 (236)
IIbPEG-b-PAA8.2TD105341.41255 (6)[38]
139444 (6)
181604 (14)
IIbPI-b-PEG8TD54n.a.1.41131 (n.a)[35]
89250 (n.a)
96353 (n.a)
21616 (n.a)
IIbPEG-b-PBLG6–7CP15751.41180 (90)[30]
8–106320.195 (20)
8–10732590 (19)
8–108729.7105 30)
10–1510920500 (126)
IIbGCPQ4.8TD140n.a152 (79)[24]
IIbPEG-b-PLGA7TD73411.5333 (n.a)[25]
IIbPTEA-b-PAM6.3CP11120.4739 (2)[34]
7032274 (3)
1701502162 (9)
IIbPEI-b-PCL-b-PEG4TD60n.a1.4120 (n.a)[28]
413056 (n.a)
417072 (n.a)
7.545100 (n.a)
7.580200 (n.a)
7.5130175 (n.a)
8.745115 (n.a)
8.780235 (n.a)
8.718070 (n.a)
9.85550 (n.a)
9.8120375 (n.a)
9.8190350 (n.a)
11.850200 (n.a)
11.8100420 (n.a)
11.8220100 (n.a)
IIclauric acid-irinotecan prodrug20TD (Sigma-Aldrich)11767189 (n.a)[33]
IIdsilica7CP24257179 (n.a)[20]
4127779 (n.a)
26421395 (n.a)
IIdsilica6.1TD16050.47148 (510)[27]
1207.4164 (607)
3135.9326 (1917)
IIIPAAn.aPolyol791001.41405 (n.a)[75]
122508 (n.a)
IIIPAA7.5Polyol151000.47247 (n.a)[49]
930340 (n.a)
11.650364 (n.a)
19.7100100 (n.a)
IIIPAA15.6Polyol371000.23361 (3.5)[74]
1238.5365 (3.4)
13.544.3319 (3.1)
1127289 (3.1)
1 molar ratio; 2 Number of nanoparticles per assembly. Abbreviation: IO: iron oxide, NF: nanoflowers, TD: thermal decomposition, CP: co-precipitation, PAA: poly(acrylic acid), PS: polystyrene, PTMC: poly(trimethylene carbonate), PGA: poly(L-glutamic acid), PEG: poly(ethylene oxide) or polyethylene glycol, PCL: poly(ε-caprolactone), PLGA: poly(lactic-co-glycolic acid), PEI: poly(ethylene imine), PI: polyisoprene, PTEA: poly(trimethylammonium ethylacrylate methyl sulfate), PAM: poly(acrylamide), PBLG: poly(γ-benzyl-L-glutamate), PR: polyrotaxane, GCPQ: N-palmitoyl-N-monomethyl-N-N-dimethyl-N-N-N-trimethyl-6-O-glycolchitosan, EPC: egg L-α-phosphatidylcholine, DSPE-PEG-methoxy 2000: 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (ammonium salt), DOPG: 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)] (sodium salt), DOPC: 1,2-dioleoyl-sn-glycero-3-phosphocholine, SDS: Sodium dodecyl sulfate.

Share and Cite

MDPI and ACS Style

Lartigue, L.; Coupeau, M.; Lesault, M. Luminophore and Magnetic Multicore Nanoassemblies for Dual-Mode MRI and Fluorescence Imaging. Nanomaterials 2020, 10, 28. https://doi.org/10.3390/nano10010028

AMA Style

Lartigue L, Coupeau M, Lesault M. Luminophore and Magnetic Multicore Nanoassemblies for Dual-Mode MRI and Fluorescence Imaging. Nanomaterials. 2020; 10(1):28. https://doi.org/10.3390/nano10010028

Chicago/Turabian Style

Lartigue, Lénaïc, Marina Coupeau, and Mélanie Lesault. 2020. "Luminophore and Magnetic Multicore Nanoassemblies for Dual-Mode MRI and Fluorescence Imaging" Nanomaterials 10, no. 1: 28. https://doi.org/10.3390/nano10010028

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop