Parasites under the Spotlight: Applications of Vibrational Spectroscopy

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Cite This: Chem. Rev. 2018, 118, 5330−5358

Parasites under the Spotlight: Applications of Vibrational Spectroscopy to Malaria Research David Perez-Guaita,† Katarzyna M. Marzec,‡,§ Andrew Hudson,⊥ Corey Evans,⊥ Tatyana Chernenko,∥ Christian Matthaü s,¶,□ Milos Miljkovic,# Max Diem,○ Philip Heraud,† Jack S. Richards,△,▽,⬡ Dean Andrew,△ David A. Anderson,△ Christian Doerig,■ Jose Garcia-Bustos,■ Don McNaughton,† and Bayden R. Wood*,† †

Centre for Biospectroscopy, School of Chemistry, Monash University, Clayton, Victoria 3800, Australia Jagiellonian Centre for Experimental Therapeutics (JCET), Jagiellonian University, Bobrzyńskiego 14, Kraków 30-348, Poland § Center for Medical Genomics (OMICRON), Jagiellonian University, Kopernika 7C, Krakow 31-034, Poland ⊥ Department of Chemistry, University of Leicester, University Road, Leicester LE1 7RH, United Kingdom ∥ Becton Dickinson and Company, 2350 Qume Drive, San Jose, California 95131, United States ¶ Leibniz Institute of Photonic Technology, Albert Einstein Straße 9, Jena 07745, Germany □ Institute of Physical Chemistry and Abbe School of Photonics, Friedrich Schiller University, Helmholtz Weg 4, Jena 07743, Germany # Department of Mechanical Engineering, Tufts University, 200 Boston Avenue, Medford, Massachusetts 02155, United States ○ Laboratory for Spectral Diagnosis (LSpD), Department of Chemistry and Chemical Biology, Northeastern University, 316 Hurtig Hall, 360 Huntington Avenue, Boston, Massachusetts 02155, United States △ Centre for Biomedical Research, Burnet Institute, Melbourne, Victoria 3004, Australia ▽ Department of Microbiology, Monash University, Clayton, Victoria 3800, Australia ⬡ Department of Medicine, University of Melbourne, Parkville, Victoria 3050, Australia ■ Department of Microbiology and the Biomedical Discovery Institute, Faculty of Medicine, Nursing and Health Sciences, Monash University, Wellington Road, Clayton, Victoria 3800, Australia

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‡

ABSTRACT: New technologies to diagnose malaria at high sensitivity and specificity are urgently needed in the developing world where the disease continues to pose a huge burden on society. Infrared and Raman spectroscopy-based diagnostic methods have a number of advantages compared with other diagnostic tests currently on the market. These include high sensitivity and specificity for detecting low levels of parasitemia along with ease of use and portability. Here, we review the application of vibrational spectroscopic techniques for monitoring and detecting malaria infection. We discuss the role of vibrational (infrared and Raman) spectroscopy in understanding the processes of parasite biology and its application to the study of interactions with antimalarial drugs. The distinct molecular phenotype that characterizes malaria infection and the high sensitivity enabling detection of low parasite densities provides a genuine opportunity for vibrational spectroscopy to become a front-line tool in the elimination of this deadly disease and provide molecular insights into the chemistry of this unique organism.

CONTENTS Introduction 1.1. Malaria 1.2. Plasmodium Parasite 1.3. Importance of Blood Stage in Diagnostics and Treatment 1.4. Current Methods for Malaria Diagnostics 1.5. Vibrational Spectroscopy Alternative 2. Instrumentation 2.1. Raman Spectroscopy 2.1.1. Single Point Measurements © 2018 American Chemical Society

2.1.2. Raman Mapping/Imaging 2.1.3. Surface Enhanced Raman Scattering and Tip Enhanced Raman Scattering 2.2. Fourier Transform Infrared Spectroscopy 3. Applications of Raman Spectroscopy in Malaria Research 3.1. Band Notation Scheme for Heme Molecules

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Received: November 2, 2017 Published: April 20, 2018 5330

DOI: 10.1021/acs.chemrev.7b00661 Chem. Rev. 2018, 118, 5330−5358

Chemical Reviews 3.2. Resonance Enhanced Raman Spectroscopy of Heme 3.3. Raman Spectroscopy of Control and Malaria Infected Red Blood Cells 3.4. Enhanced Overtones from Green Laser Light 3.5. Mechanism of Hz Formation Studied with Raman Spectroscopy 3.6. Raman Studies on the Structure and Effect of Antimalarial Drugs 3.7. 3D Visualization of Infected Red Blood Cells 4. Diagnosis of Malaria Using Raman Methods 4.1. Use of Raman Microscopes on the Detection of Hz in RBC 4.2. Applications of SERS, SERRS, and SORS for Plasmodium Detection 5. FTIR Spectroscopy as a Diagnostic Tool for Malaria 5.1. Assignment of FTIR Spectrum from Malaria Infected Cells 5.2. Synchrotron FTIR 5.3. Focal Plane Array (FPA) Imaging Using a Conventional Source 5.4. ATR-FTIR 6. Conclusions and Outlook Author Information Corresponding Author ORCID Notes Biographies Acknowledgments References

Review

Only female Anopheles mosquitoes transmit the malaria parasite. Sporozoites in the insect’s saliva are deposited during a bite and migrate from the skin to the liver, where the parasite numbers increase while hidden from the immune system. They then emerge, infect RBCs, and reproduce, generating new parasite progeny every 48−72 h depending on the infecting Plasmodium species. Highly synchronous parasite emergence induces the cyclical febrile paroxysms typical of the disease. P. falciparum infected RBCs tend to sequester in the deep vasculature and thus avoid being cleared from the host in the spleen. This can however result in circulatory blockage and inflammatory reactions that lead to organ failure and cerebral malaria, usually fatal if not treated quickly with efficacious drugs,5 which explains why P. falciparum is the agent of severe malaria.

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1.2. Plasmodium Parasite

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The life cycle of Plasmodium species is complex and involves a vertebrate host and an arthropod vector for all five species that infect humans, with the vector being a mosquito of the genus Anopheles. Infection of the human host begins with the injection of Plasmodium spp. sporozoites into the dermis by an infected mosquito during a blood meal. The motile sporozoites enter a blood vessel at the site of inoculation and reach the liver through the bloodstream. There, they invade hepatocytes, beginning with an asexual replication process called exoerythrocytic schizogony (as mentioned above, P. vivax and P. ovale are also able to established a latent infection in hepatocytes, in the form of dormant “hypnozoites”, which can be reactivated to a productive blood infection months or years after the initial episode). The ensuing liver schizont produces tens of thousands of merozoites, which are released into the bloodstream and are primed to invade RBCs. Each infected cell ultimately generates about 20 new merozoites that are released into the bloodstream to infect new RBCs within seconds. Circulating antibodies against merozoite antigens have to neutralize the pathogen within these few seconds. The iterative synchronous infection and lysis of RBCs is the cause of malaria pathology. Parasite waste products elicit a strong inflammatory response, including high fever, lactic acidosis, and anemia. When inflammation occurs in the brain following adherence of P. falciparum-infected RBCs to the vasculature, the result is cerebral malaria, which has a fatal outcome if not promptly treated.

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INTRODUCTION 1.1. Malaria

Over 3.4 billion people are at risk of malaria, making it the most devastating vector-borne disease on the planet. Despite a considerable drop in mortality over recent decades, from up to 3 million deaths annually to less than 500 000 in 2016, malaria remains one of the world’s leading causes of childhood death.1 This improvement is the result of a combination of transmission control measures such as massive deployment of insecticide-treated bed nets and the implementation of artemisinin combination therapies (ACT). Unfortunately, the fight against malaria is far from over, and the spread of insecticide-resistant mosquitoes and the emergence of parasites with reduced susceptibility to artemisinin derivatives is cause for serious concern. Here, we give a short introduction to malaria research, and more in-depth reviews on the disease can be found in recent reviews.2−4 Human malaria is caused by infection with parasitic protists of the genus Plasmodium, five species of which infect humans (P. falciparum, P. vivax, P. ovale, P. malariae, and P. knowlesi). P. falciparum is the most virulent species and causes the majority of lethal malaria cases (mostly in sub-Saharan Africa), a feature associated with the ability of the infected red blood cells (RBCs) to adhere to endothelial cells in the vasculature. P. vivax, however, is responsible for a heavy morbidity burden and has the capacity (together with P. ovale) to establish a latent asymptomatic infection inside hepatocytes, complicating control programs.

1.3. Importance of Blood Stage in Diagnostics and Treatment

Each of the five Plasmodium species known to parasitise humans has its preferred anopheline vectors and hence geographical distribution. There are a range of malaria parasites adapted to all terrestrial vertebrates, and those able to grow in humans were originally probably zoonotic parasites, passing back and forth between primates and humans, with P. knowlesi still currently transmitted from reservoirs in SE Asian monkeys. P. falciparum is believed to have made a relatively recent speciation jump from gorillas.6 It may still be adapting to humans, an ongoing process that perhaps contributes to the reasons why it causes 99% of malaria deaths.1. The next most prevalent species, P. vivax and P. ovale, have similar modes of infection in that some of the parasites reaching the liver do not immediately enter the division cycle, establishing instead a latent infection reservoir that produces recurrent blood infections if not specifically treated. This has important 5331


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in peripheral blood is the ring, making it an ideal target for a diagnostic test using blood acquired from a patient. The other noncirculating stages such as trophozoites and schizonts present much more drastic phenotypic features and are more easily distinguished by molecular methods from uninfected RBCs. Both are mature stages which contain a considerable amount of hemozoin (Hz), lipids, and DNA, but they are rarely found in blood obtained from standard peripheral blood sampling.

implications for diagnostics and therapy as well as for ongoing disease elimination and eradication efforts. Most existing treatments target the blood borne stages of the disease illustrated in Figure 1. This makes sense inasmuch as

1.4. Current Methods for Malaria Diagnostics

Early detection and treatment of malaria is critical in preventing morbidity and reducing mortality. Point-of-care (POC) testing for malaria infection is generally reliant on antigen capture, immune-chromatographic-based rapid diagnostic tests (RDTs), which have been extremely effective in detecting relatively highdensity parasite infections typically found in symptomatic individuals. Malaria elimination strategies require diagnostic tools that can detect individuals with low-density infection.8,9 This is especially the case for P. vivax but also asymptomatic carriers of P. falciparum.10 Unfortunately, current RDTs lack the sensitivity and specificity to detect parasite infection at these low densities.11,12 Light microscopy also suffers from suboptimal sensitivity, requires highly trained technicians, and has proven hard to implement in many resource-poor regions. Highly sensitive methods like polymerase chain reaction (PCR) detection, which is used primarily as a research tool, are too expensive and technically challenging for widespread implementation.

Figure 1. Blood stages of the Plasmodium parasite species in blood. Reproduced with permission from ref 7. Copyright 2014 American Chemical Society.

these stages are responsible for the observed pathology, but disease eradication will require targeting the liver reservoirs and probably also the forms able to infect mosquitoes. The latter forms are sexually differentiated and largely metabolically inactive parasites called gametocytes. They are able to survive digestion of the blood meal in the insect midgut and so available to complete the sexual part of the parasite’s life cycle in the mosquito. Mature gametocytes are difficult to target pharmacologically given their reduced metabolic activity, but immature forms could be more easily targeted, and active insect stages can also in theory be killed with drugs taken up by the mosquito with the blood meal. Such drugs would not be directly beneficial to the infected person, but they would protect the community by curtailing disease transmission. Gametocytes are produced at different times following exit from the liver with the specific times dependent on the particular Plasmodium species. This gametocyte production sometimes occurs even before symptoms develop. Additionally, in high-transmission areas, partial immunity allows people to harbor parasites without suffering disease symptoms. The production of infective gametocytes without overt disease symptoms makes prompt diagnosis and effective treatment crucial for public health interventions. The initial stage after invasion of the RBC infection and the most common stage found in peripheral blood of P. falciparuminfected patients is termed the ring stage, from its ring-like appearance in Giemsa-stained blood films. More mature stages are sequestered in the deep vasculature because of parasiteencoded adhesins expressed at the surface of the infected RBC and are therefore usually not detected in peripheral blood. The ring stage is metabolically quiescent and therefore generally resistant to most antimalarial drugs. Rings transition into the trophozoite stage approximately 12 h postinvasion, and during a further 12−14 h, the trophozoite actively grows at the expense of the host cell and serum components. At 26−28 h postinvasion, the first nuclear division defines the entry into the schizont stage, characterized by an increasing number of nuclei. The active growth and replication that characterizes trophozoites and schizonts make these stages the most sensitive to antimalarial drugs. Starting about 42 h postinvasion, daughter merozoites, assembled after the last round of nuclear divisions, burst out of the remnant RBC. The most common stage found

1.5. Vibrational Spectroscopy Alternative

New modalities that are rapid, cheap, and highly sensitive and that can diagnose multiple Plasmodium species simultaneously using small sample volumes of blood are urgently required. Infrared and Raman spectroscopy are molecular-based approaches that rely on the detection of macromolecules13,14 specific to the sample under analysis. These label-free and nondestructive vibrational techniques, Fourier transform infrared (FTIR), and Raman spectroscopy enable the collection of characteristic spectra, which are unique for each molecular system and can serve to identify compounds unambiguously. FTIR spectroscopy is an absorption-based technique that detects vibrational transitions in molecules that have an intrinsic change in dipole moment during the oscillations of the nuclei in the molecules. The technique is eminently suitable for the analysis of macromolecules like proteins, nucleic acids, carbohydrates, and lipids15 and hence has enormous potential diagnostic capability in distinguishing Plasmodium infected red blood cells (iRBCs) from uninfected cells. Raman spectroscopy, on the other hand, is a scattering technique where a dipole moment must be induced by the electric field of the incident laser to create energy shifts that reflect changes in the vibrational energy levels of the molecule.16 Raman spectroscopy, similarly to IR, depends on molecular vibrations, and observed Raman shifts occur in the same energy range as IR absorption.17 Molecules that tend to be good Raman scatterers are often very symmetric, and the signal intensity also increases for chromophoric or conjugated systems.18 Hemozoin (also known as malaria pigment) is an ideal Raman scatterer because of the high symmetry of the dimeric heme units and the chromophoric nature of protoporphyrin IX. Both Raman and infrared microscopic techniques are diffraction limited, and therefore, their lateral resolutions are dependent on the wavelength of light used to excite the 5332


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Figure 2. Comparison of the different approaches for obtaining Raman spectra of cells and parasites showing single point mapping, Raman microscopy, SERS, and TERS and their relative features. The theoretical diffraction limits shown for each technique were calculated from the Rayleigh criterion (Δxy ≈ 0.61λ/NA; Δz = λ/NA2) for the excitation range 488 to 1064 nm and using objectives with NA values from 1.2 to 0.5.

sample.19 Thus, the lateral resolution of Raman microscopy, which uses visible and near-infrared light (400−1000 nm) for excitation, is approximately an order of magnitude greater than infrared microscopy, which utilizes mid infrared light (2.5−15 μm). Furthermore, water has a high molar extinction coefficient but a very small Raman cross-section and hence is a weak Raman scatterer. For this reason, Raman spectroscopy remains the preferred technique for live cell analysis. This review focuses on the application of both Raman and infrared spectroscopy to malaria research, including instrumentation, spectral interpretation, and the diagnostic and monitoring capability of these new modalities.

scattering) or a different energy (inelastic scattering, Raman scattering).27 We may distinguish two types of Raman scattering: Stokes scattering where the scattered photon has reduced energy, and anti-Stokes scattering, where the scattered photon has higher energy. The Stokes Raman spectrum results from the original molecule being in the ground vibrational state and the anti-Stokes results from molecules in excited vibrational states. Hence, the Stokes spectrum is the most intense and is recorded in most available spectrometers and confocal microscopes.33 The resultant spectrum is used to identify functional groups in the sample as well as study their distribution in 2D (Raman imaging/mapping) and 3D (Raman imaging/mapping for different planes/stacks). The lateral and depth resolutions of confocal Raman microscopy depends on the laser wavelength (λ) used as well as on the numerical aperture (NA) of the objective and the size of the confocal aperture. The theoretical diffraction limits can be calculated from the Rayleigh criterion (Δxy ≈ 0.61λ/NA, Δz = λ/NA2) but are rarely achieved in practice because of the limitations of the confocal pinhole size.18 Because inelastic light scattering has a very weak effect, it is essential in many studies to find ways to enhance the Raman signal. Surface Enhanced Raman Scattering (SERS) is one method of Raman signal enhancement through the interaction of a molecule with a nanostructured metal/metalloid substrate.34 The high values of enhancement of this plasmon-assisted scattering depend on the type of molecule as well as on the quality of the metallic/ metalloid substrates (e.g., nanoparticles suspensions, electrochemically roughened electrodes). Tip-enhanced Raman Scattering (TERS) can be viewed as a type of SERS technique where the contact between molecule and metallic substrate is produced by a metallized tip controlled by an atomic force microscope (AFM) allowing also TERS imaging.35 SERS and TERS provide lateral and depth resolution below the theoretical diffraction limits and allow for single-molecule detection.36,37 Additionally, for both SERS and TERS, when the excitation energy matches the energy of the allowed molecular electronic transition (as in the case of resonance Raman spectroscopy), Surface-enhanced resonance Raman scattering

2. INSTRUMENTATION 2.1. Raman Spectroscopy

Figure 2 shows schematic representations of the different approaches employed for Raman measurements using a microscope platform or a fiber optic system, which is the most common form of modern laboratory-based instrumentation. Each acquisition mode offers different possibilities and challenges in terms of sample preparation, spectral acquisition, and analysis, and these are critically evaluated in the following sections. The discovery,20 theory,16,21 and instrumentation22,23 of Raman scattering have been reported in many monographs. The latest works are mainly focused on the application of different Raman techniques to chemical analysis24−26 and the field of biomedical studies27,28 or life sciences.29,30 Because the scope of this review is not to provide an overview of Raman microscopy, which can be found elsewhere,31 here, we present only an introduction to the Raman scattering phenomenon. Raman scattering is a process where a photon of the light interacts with a molecule and induces a transition of the molecule to a virtual energy state above the original state of the molecule. If the photon energy is such that an excited electronic state of the molecule is accessed, then the result is resonance Raman scattering.32 The molecule can relax back on a short time scale to its initial ground state with the resultant photon having either the same energy (elastic scattering, Rayleigh 5333


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laser coupled to the WITec Raman microscope to generate 3D images of Plasmodium iRBCs. Raman imaging differs from Raman mapping in that a filter is used in the former to select a small range of the Raman scattered light corresponding to a Raman band or bands. The scattered light over a large sample area outlining the object of interest is then focused on the CCD, and an intensity map derived from the CCD array then generates an image of the field of view.42 This approach was used to analyze malariainfected erythrocytes at the late trophozoite stage using a 780 nm diode laser with a 120 s laser exposure time.43 The image was collected by first calibrating the filter to the position of a characteristic Hz band (ν4 = 1374 cm−1) by recording a filter spectrum of the Hz over the 1800−200 cm−1 range. Images were recorded by defocusing the laser approximately 15% to encapsulate the whole cell, and the scattered light was transmitted through a polarizer. A background image was also recorded around 1800 cm−1, a spectral range containing no spectral bands, and this image was subtracted from the ν4 image to minimize the effects of the background and to remove inconsistencies within the laser spot. Using this approach, we demonstrated that it was possible to detect Hz from a mature trophozoite in a single functional erythrocyte.43 2.1.3. Surface Enhanced Raman Scattering and Tip Enhanced Raman Scattering. Since the discovery of enhanced Raman signals from pyridine on a silver electrode in the 1970s,44 the development of SERS for the detection and investigation of Raman active compounds has been constantly evolving.45 In a SERS experiment, the presence of metal (i.e., Au or Ag) nanostructures close to a target molecule enhances some Raman modes of the compound either through an electromagnetic plasmon effect and/or by a chemical effect, where an electronic charge transfer transition is generated by the interaction of the molecule with the nanostructure.46 This property can be used for improving the limit of detection of an analyte by many orders of magnitude47 compared with normal Raman spectroscopy. Conventional Raman spectrophotometers can be used for measuring SERS spectra, making this technique easy to implement, cost-effective, and potentially suitable for POC diagnosis especially with the incorporation of Raman optic fiber probes.48,49 An example is the enhancement of the signal of porphyrins in the so-called fiber enhanced Raman spectroscopy (FERS),50 where the molecules are excited and the Raman signal collected along the path length of the hollow core of a fiber. In the case of malaria diagnosis, the Ramanscattered signal from functional groups in hemoglobin (Hb) and Hz can be enhanced 51 provided that the metal nanostructures are close in contact with the Hz crystals. AFM tips can be coated with metal nanoparticles that serve as SERS active particles for molecules in the proximity of the tip.35 This is a relatively new technique which combines the spatial resolution of AFM with the molecular selectivity provided by Raman spectroscopy. TERS microscopes enable the imaging of features on the nanoscale, providing information about the molecular structure.25 However, the low penetration depth of this technique limits its application in biological sciences to the surface of cells, viruses, and large biomolecules.52 Moreover, tip reproducibility, molecular orientation, and localized heating of the sample area make band assignment and spectral interpretation difficult.

(SERRS) and tip-enhanced resonance Raman spectroscopy (TERRS) are achieved. 2.1.1. Single Point Measurements. The simplest approach to obtain a Raman spectrum of a living single RBC is to focus the laser beam directly onto the cell using a visible microscope and water immersion objective.38 The Raman signal is back scattered through the microscope objective to a spectrophotometer for spectral acquisition using a diffraction grating and CCD detector. This configuration is widely used in biological/biomedical research laboratories because of the high spatial resolution that enables mapping of subcellular structures. Raster mapping can be used to collect 2D Raman maps, although this is generally a time-consuming process. The acquisition time can be reduced by a method where the beam is diffused into a line rather than a point focus and lines of spectra can be collected, i.e. Raman Streamline (Renishaw). This is also advisable when dealing with photosensitive cells because it reduces the laser power on a given area. Alternatively, a new range of spectrometers incorporating fast acquisition spectrophotometers and piezoelectric motorized stages can be utilized, as described in the following section. A number of important parameters must be controlled when performing Raman measurements on single living RBCs. Laser power at the sample and laser exposure time are two critical parameters that need to be optimized in these types of experiments. High laser power can lead to photodenaturation, aggregation, and spectral baseline effects,39 while low laser power can result in a poor signal-to-noise ratio unless the acquisition time is long and hence prohibitive for mapping. The laser should be shuttered to eliminate the unnecessary exposure to radiation between successive measurements. To control the physiological environment of cells, a temperature-controlled stage that can accommodate a Petri dish is required. In this configuration, the cells can be examined directly in phosphate buffered saline using a water immersion objective.39 Coating the Petri dish with a reflective coating such as aluminum is advisable to avoid Raman signals from the material of the Petri dish. Although the reflective coating can affect the confocality, it improves the signal-to-noise ratio by also capturing reflected Raman scattered light.40 Cells are usually fixed to the Petri dish via a surface coating of poly L-lysine.41 2.1.2. Raman Mapping/Imaging. A range of Raman systems are available that utilize fiber optic delivery to obtain true confocal Raman microspectroscopic maps. These can also utilize ultrafast electron multiplying charge-coupled device (EMCCD) detectors to enable more rapid spectral acquisition. These systems have revolutionized confocal Raman mapping. The RBC mapping work at Monash University has been performed mainly on a WITec confocal CRM alpha 300 Raman microscope. The spectrometer is equipped with an air-cooled solid−state laser operating at 532 nm and an ultrafast EMCCD detector thermos-electrically cooled to −60 °C. The laser is coupled to the microscope via an optical fiber with a diameter of 50 μm (other diameters are available). The scattered radiation is focused onto a multimode fiber (50 μm diameter) and a single monochromator (focal length of 300 mm and an aperture ratio equal to f/4). The spectral resolution is equal to about 3 cm−1 (600/mm grating). The lateral spatial resolution achieved using a water immersion objective (60X, NA = 0.6) and 532 nm excitation is approximately 540 nm. An alternative imaging configuration was used for the 3D image presented below (Section 3.1). This was obtained using a 633 nm HeNe 5334


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Figure 3. Schematics and features of the different approaches used for obtaining the infrared spectrum of RBCs showing synchrotron radiation, focal plane array (FPA), attenuated total reflectance (ATR).

2.2. Fourier Transform Infrared Spectroscopy

straight sections. As the electrons are deflected through the magnetic field created by the magnets, they emit electromagnetic radiation, so that at each bending magnet, two beams of synchrotron light, termed edge radiation and bendingmagnet radiation, are produced. Both beams contain infrared radiation, which can be used separately or can be combined to increase the flux. In the case of an IR microscopy beamline, the IR light is focused onto the sample and single point maps with a high signal-to-noise ratio, and a spatial resolution close to the diffraction limit of the infrared light can be obtained. The use of bright synchrotron light is especially useful for hydrated cells, which normally require a powerful IR source due to the high extinction coefficient of water in the mid-IR region and the fact that the light needs to pass through two chamber windows. More recently, synchrotron facilities have been working on expanding the beam or irradiating samples with multiple beams to make use of focal plane array imaging detectors.58 This new configuration allows the acquisition of images as opposed to single point maps, thus reducing the measurement time. The focal plane array (FPA) system is normally operated with a classical infrared thermal source such as a hot wire or globar.59 Nowadays FPA detectors contain an array of up to 128 × 128 MCT detectors, which enable the simultaneous acquisition of hyperspectral images of 16 384 pixels.60 Alternatively, smaller 16 pixel detectors such as in the PerkinElmer Spotlight can be used to rapidly raster scan samples. One of the main challenges of using a conventional IR microscope system, besides the low signal-to-noise ratio from using a thermal source, is the low lateral resolution, which restricts the imaging capability to cells and cellular components with a size of approximately 5 μm. However, high magnification elements and objectives can overcome this problem by oversampling the images while approaching the diffraction limit. In a recent article, high resolution FTIR images were obtained using a state-of-the art FPA microscope equipped with a high magnification element (4×) and a new 25× Cassegrain optic with a high NA = 0.81, which enabled the acquisition of images with a pixel size of 0.66 μm and a lateral resolution of 1.4 μm at 3750 cm−1.61

The use of infrared spectroscopy for research and diagnosis in the clinical fields has been addressed in several reviews and monographs.15,53−55 Infrared spectra of RBCs can be acquired with a wide range of instruments ranging from expensive and sophisticated FTIR imaging microscopes to portable spectrometers.56 The spectrum is obtained by placing the sample directly in the light path between an IR light source and a detector. An interferometer allows for the modulation of different wavelengths at different rates, which simultaneously impinge on the sample with the transmitted or reflected light focused to a detector. A Fourier transform is applied to transfer the resultant interferogram signal from the optical path difference (cm) domain into the wavenumber domain (cm−1), and a transmission spectrum is obtained by ratioing the output against a background spectrum, usually air or solvent from the sample spectrum. An absorbance spectrum, directly related to sample concentration, is obtained by taking the log10 of the transmittance spectrum T = P/P0, where P is the transmitted radiant power and P0 is the incident radiant power. The absorbance of the sample is therefore A = log10 P0/P. Different source configurations and detectors can be selected for obtaining the spectra depending on the spatial resolution and signal-to-noise ratio required. Figure 3 depicts three types of FTIR configurations reviewed here. Although the diffraction limit of a benchtop FTIR microscope system allows for spectroscopy of a single RBC, diameter ca. 7 μm, the microscope aperture limits the photon flux and hence the signal/noise. The signal-to-noise ratio can be improved with these small apertures if a bright synchrotron source is utilized. Synchrotron FTIR commonly uses a single point IR microscope but utilizes highly collimated and often apertureless synchrotron radiation as the light source.57 A bright-collimated IR beam is obtained by accelerating electrons around a synchrotron ring. Electrons are generated by an electron gun and accelerated to 99.9997% of the speed of light by a linear accelerator. The electrons are then transferred to a booster ring, where their energy is increased before being transferred to an outer storage ring. The electrons are circulated around the storage ring by a series of magnets separated by 5335


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Figure 4. (a) Chemical structure of β-hematin. (b) Structural diagram of the protoporphyrin IX with standard atom labeling.67

in the D4h symmetry point group.68 When the hydrogen atoms are treated as point masses, the 37 atom molecule has 71 inplane modes (2N − 3), giving the following irreducible representation.

In cases where high lateral resolution is not required, ATRFTIR spectrophotometers can acquire a spectrum from a sample in a simple and cost-effective way. This technique is the standard methodology in many laboratories for functional group identification and has been widely employed in the analysis of multiple cells and tissues.62−64 The sample is placed on an IR-transparent crystal with a high refractive index. The IR beam is passed through the crystal at an angle that results in the beam being totally internally reflected. An evanescent wave penetrates into the sample in contact with the crystal. The penetration depth of the evanescent wave is wavelength dependent, typically between 0.5 and 5 μm for IR and also depends on the angle of incidence and the refractive index of both crystal and sample.65 Nowadays, infrared instrumentation can be miniaturized, and hand-held spectrophotometers are available for around $10 000. In addition, Si-based disposable ATR crystals have been developed66 and are already being commercialized by IRUBIS GmbH (Munich, Germany). With a standard low cost, portable spectrometer and an ATR accessory, a high-quality spectrum of RBCs can be obtained in less than a minute, making the technique an ideal POC device for malaria diagnosis.

Γin‐plane = 9A1g + 8A 2g + 9B1g + 9B2g + 18E u

The “gerade” modes except for A2g are only Raman active, while the “ungerade” modes, i.e. Eu modes, are only IR active. A2g modes are both Raman and infrared active. There are also 34 out-of-plane modes with the following irreducible representation. Γout ‐ of ‐ plane = 3A1u + 6A 2u + 5B1u + 4B2u + 8Eg

For a perfect D4h symmetric molecule only the Eg out-ofplane modes are Raman active, but in reality, because of porphyrin ruffling and other distortions, other modes become allowed. The symbol ν designates an in-plane stretching mode, while γ represents an out-of-plane mode. The subscript numbering denotes the mode number and is based on convention with the numbering starting at the high wavenumber most symmetric modes and finishing with the lowest wavenumber most nonsymmetric mode. Therefore, the inplane modes are designated as A1g (ν1 − ν9), B1g (ν10 − ν19), B2g (ν20 − ν29), A2g (ν30 − ν19), and Eu (ν40 − ν49). The basic symmetry of the vibrational modes can be determined experimentally by the depolarization ratio defined

3. APPLICATIONS OF RAMAN SPECTROSCOPY IN MALARIA RESEARCH 3.1. Band Notation Scheme for Heme Molecules

Due to the high Raman activity of both Hb and Hz, the bands of these two components dominate the spectra of RBCs infected with the Plasmodium parasite. Hb is the iron containing protein responsible for transporting oxygen in humans and can be found in two main conformations: linked to an O2 molecule (OxyHb) or free (DeoxyHb). Hz results from the catabolization of Hb in the digestive vacuole of the malaria parasite. Powder diffraction data of β-hematin (Fe(III)PPIX)2, a spectroscopically identical synthetic analogue of Hz, obtained with synchrotron radiation has shown it to be an array of heme dimers linked through reciprocal iron-carboxylate bonds to one of the propionate side chains, and these dimers in turn form chains linked by hydrogen bonds48 (Figure 4a). The central prosthetic group in Hb and Hz is known as iron protoporphyrin IX and is shown in Figure 4b using standard notation67 to label the carbon atoms. It consists of a porphyrin group with an iron center that contains four pyrrole groups and four unsaturated methylene carbons. The high symmetry and chromophoric structure results in a large Raman cross-section and hence strong Raman scattering when exciting with UV/ visible/near-IR laser lines. The heme part of the molecule has a fourfold principal axis of symmetry along with a rotation/ reflection plane and a horizontal plane of symmetry, placing it

as ρ =

I I⊥

where I∥ is the intensity of the parallel component of

scattered light and I⊥ is the intensity of the perpendicular component. Totally symmetric modes of A1g symmetry, assuming the D4h point group, have ρ < 0.5, while nontotally symmetric modes of B1g, B2g, and A2g symmetry have ρ > 0.5. This is particularly useful in assigning overtone modes based on the symmetry combinations of the fundamental modes. 3.2. Resonance Enhanced Raman Spectroscopy of Heme

The strong Raman scattering by heme pigments such as Hz and Hb makes this technique ideal for studying the host−parasite interactions. For such chromophoric compounds, it is advantageous to use resonance Raman spectroscopy where the incident laser is in resonance with an electronic transition and accesses the vibronic energy levels. The intensities of Raman scattering signals can be increased by up to three orders of magnitude, leading to a much more sensitive technique. Ong et al.69,70 were the first to apply resonance Raman spectroscopy to normal and P. berghei-infected mouse erythrocytes fixed in methanol using 488 nm excitation21 and later 633 nm excitation.70 It has been demonstrated that it is possible to record high S/N Raman spectra of β-hematin at a range of different excitation 5336


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Figure 5. (a) UV−visible spectra of hemoglobin in the oxygenated and deoxygenated forms featuring the different electronic transitions. Reproduced with permission from ref 72. Copyright 2014 Wiley. (b) Absorption spectra of Hz (A) and β-hematin (B). Reproduced with permission from ref 73. Copyright 2009 Wiley.

wavelengths71 and to detect Hz within the food vacuole of a functional RBC infected within a P. falciparum trophozoite.43 The electronic absorption spectrum of Hb and Hz (see Figure 5) is dominated by three strong bands resulting from combinations of π ← π* electronic transitions and known as the Soret band (∼400 nm), the Qv band (∼520 nm), and Q0 band (∼550 nm). Excitation into the Soret band results in strong scattering of the totally symmetric A1g modes through a Type A or Franck−Condon mechanism, whereby displacement of the normal coordinate during nuclei oscillation results in the overlap of the ground and excited state vibrational wave functions, leading to the enhancement of the totally symmetric modes. Excitation into the Q0 band at ∼550 nm results in what is known as Type B scattering, whereby the displacement of the nuclear coordinate during oscillation couples the ground and excited state electronic wave functions in a process known as vibronic coupling. This results in the enhancement of nontotally symmetric modes, including B1g, B2g, and A2g modes. Enhancement in the Qv band results in type B scattering and also another type of scattering process known as type C scattering. In type C scattering, a low frequency vibrational mode can couple two forbidden energetically similar electronic transitions, giving rise to the enhancement of nontotally symmetric modes and also providing a mechanism for the appearance of overtone modes, as will be discussed. The Raman spectra of hemin, β-hematin, and related compounds have been extensively studied using a wide range of excitation wavelengths.71,72 A study71 revealed that the Raman spectra of β-hematin obtained with near-infrared laser excitation (780 and 830 nm) showed dramatic enhancement of the out-of-plane totally symmetric A1g and breathing modes (Figure 6). This is quite unusual because totally symmetric modes of heme complexes are normally enhanced only when the excitation laser is in resonance with the Soret band at ∼400 nm. This enhancement is also observed in hemin (FePPIX-Cl), the monomeric precursor of β-hematin, but to a lesser extent. These differences were explained by considering intermolecular

excitonic interactions between porphyrinic units. The enhancement was explained by invoking the aggregated enhanced raman scattering (AERS) hypothesis originally espoused by Akins for porphyrins74−76 and cyanine dyes.77−79 In this case, the stacking of the hemes results in strong interactions through superposition of z-polarized charge-transfer electronic transitions, giving rise to both Type A (Franck−Condon) and Type B (vibronic coupling) enhancement.71 As discussed above, Type A enhancement is observed when exciting with laser wavelengths in the vicinity of the Soret band (∼400 nm) for all hemes, while Type B enhancement is observed when exciting with wavelengths in the vicinity of Qv (∼530 nm) and Q0 (∼575 nm). Thus, the appearance of dramatically enhanced totally symmetric modes in Hz and β-hematin, when using near-IR excitation wavelengths, which are well away from the electronic transitions, is indeed unusual and not predicted by conventional scattering theory. Consequently, to image hemozoin in red blood cells, it is better to use near-IR excitation wavelengths where the dramatic enhancement of the totally symmetric modes is observed. Moreover, using near-IR wavelengths reduces the energy on the cell, making these longer excitation wavelengths more conducive to live cell imaging. 3.3. Raman Spectroscopy of Control and Malaria Infected Red Blood Cells

The Raman spectra of iRBCs recorded at the trophozoite stage show strong bands from Hz for a range of excitation wavelengths, including 406, 488, 514, 568, 633, 647, 676, 780, 830, and 1064 nm.71,73 Hz bands dominate the Raman spectra of trophozoite-infected red blood cells, while in the case of control RBCs, the spectra contain strong bands from oxyhemoglobin (oxyHb). For some iRBC cells, where the Hz concentration is low, an intermediate state exists, and the spectra result from a mixture of Hz and oxyHb. Both Hz and oxyHb contain the heme prosthetic group, which provides strong resonance enhancement. Figure 7 shows the average Raman spectra in the fundamental spectral region obtained with the use of 514.5 nm laser excitation from approximately 30 5337


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Figure 7. Average Raman spectra obtained using 514.5 nm laser excitation from around 30 single functional, control RBCs (black) and 30 single iRBCs (red) measured in saline solution as described46 with the most prominent Raman wavenumber values (cm−1) for the fundamental bands for oxyHb (RBCs) and Hz observed inside iRBCs. The most prominent marker bands of Hz are marked with stars.

density marker band ν4 for Hz, observed at 1371 cm−1, corresponds to the Fe being in the ferric high spin state and has been successfully used to study the interaction between hematin and antimalarial drugs, as will be discussed.82 Although at first sight in Figure 7 bands from iRBCs and RBCs can look similar to the naked eye, chemometric routines such as principal component analysis (PCA) can be used to clearly differentiate the spectral markers of Hz. 3.4. Enhanced Overtones from Green Laser Light

An unusual case of resonance Raman scattering is the observation of enhanced overtone bands when exciting Hb inside the highly concentrated heme environment of the RBC using 514.5 nm excitation (see Figure 8). In this example, the overtone bands (4000−2600 cm−1) were even more intense than the fundamentals (1800−200 cm−1). This enhancement was observed only for excitation using wavelengths in the Qv band and was not observed in solutions of Hb. The enhancement of overtones was also observed in crystals of other hemes, including Hz, hemin, and hematin, and was shown to improve the diagnostic capability of Raman spectroscopy when both the fundamental and overtone regions were included in the partial least squares discriminant analysis (PLS-DA) model.72 The appearance of overtones in resonance Raman spectra of hemes is predicted by theory invoking Cterm scattering. Unlike B-term scattering, which involves vibronic coupling of a vibrational mode in the excited state, C-term scattering results when a vibrational mode couples two forbidden electronic transitions in the excited state. However, this does not explain the enormous enhancement of overtones and combination bands when exciting between 500 and 540 nm in the vibronic Qv band. A proposed hypothesis suggests that this unusual enhancement results from excitonic coupling between linked porphyrin moieties in the extended porphyrin array, which enabled the investigation of Hz within its natural environment of functional RBCs.43 The nonfundamental region from Hz is characterized mainly by 2ν10, 2ν19, 2ν37, 2ν3, and 2ν4,

Figure 6. Raman spectra of β-hematin recorded using a variety of excitation wavelengths. The asterisks in the 830 nm spectrum highlight bands that appear dramatically enhanced. Reprinted with permission from ref 71. Copyright 2004 American Chemical Society.

single functional, control RBCs and 30 single iRBCs measured in saline solution as described previously.72 During the measurements, the beam was slightly defocused onto the RBC to prevent localized heating, and the power of the laser at the sample position was ≤0.5 mW, which prevented photo/ thermal degradation and the formation of heme aggregates inside the RBCs.39,80 The Raman spectrum (see Figure 7) of control RBCs is dominated by oxyHb with the most characteristic bands in the fundamental region observed at 1639, 1605, 1587, 1547, 1473, 1431, and 1376/1357 cm−1, which are assigned to ν10, ν19, ν37, ν11, ν3, ν28, and ν4 porphyrin stretching modes, respectively (see Table 1). On the other hand, the Raman spectrum of the iRBCs in the region of the porphyrin skeletal vibrations of the heme macrocycle (1600− 1500 cm−1) is characterized by strong ν10 and ν19 modes, which are shifted to 1629 and 1570 cm−1, respectively. Moreover, the appearance of the Hz in iRBCs causes a shift of ν3 and ν4 to 1490 and 1371 cm−1, respectively. The most important marker band of Hz is the band at 1570 cm−1 assigned to the ν19 mode. This band was found to be sensitive to the interaction and binding affinity between Hz and antimalarial drugs.81 The π 5338


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Table 1. Main Raman Band Assignments of the RBCs and P. falciparum Infected RBCs Obtained Using the 514.5 nm Laser Line Showing the Resonance Raman Wavenumber Values (cm−1) for the Most Prominent Fundamental Bands, Overtones, and Combination Modes for oxyHb (RBCs) and Hz (Marked with *) Observed in iRBCs with Assignments and Symmetry Terms Assuming an Idealized D4h Symmetrya,51,72,81,83 fundamental bands obs/cm

a

−1

overtones and combination bands

assignment

symmetry

1639 1629* 1605 1587 1590 sh* 1570* 1547 1490* 1473

ν10 ν10 ν19 ν37 ν37 ν19/ ν2 ν11 ν3 ν3

B1g B1g A2g Eu Eu A2g A2g A1g A1g

local coordinate ν(CaCm)asym ν(CaCm)asym ν(CaCm)asym ν(CaCm)asym ν(CaCm)asym ν(CaCm)asym ν(CbCb) ν(CaCm)sym ν(CaCm)sym

1431/1431*

ν28

B2g

ν(CaCb)sym

1371* 1376 1357

ν4 ν4 ν4

A1g A1g A1g

ν(half-ring)sym ν(half-ring)sym ν(half-ring)sym

obs/cm

−1

assignment

symmetry

3277m 3258m 3200 br,sh 3172 s,br 3180 s,br 3140 br,sh 3100 s,br 2980−2930 br,sh 2980−2930 br,sh 2890 vs

2ν10 2ν10 2ν19 2ν37 2ν37 2ν19/ ν2 2ν11 2ν3 2ν3 (1547 + 1339) ν11 + ν41

A1g A1g A1g A1g A1g A1g A1g A1g A1g A1g

2855/2874 br,m,sh 2742 w

ν3 + ν4 2ν4

A1g A1g

2720 w

2ν4

A1g

ν, stretching; δ, bending; sym, symmetric; asym, asymmetric; vs, very strong; s, strong; m, medium; w, weak; br, broad; sh, shoulder.

which are incorporated into the parasite’s proteins and used for energy metabolism. Catabolism of Hb produces a large amount of free heme, which is thought to be toxic to the parasite, resulting in lipid oxidation85 and disruption of cell membranes,86 but other explanations have been suggested.87 To reduce the toxic effects of free heme, the parasite has evolved a detoxification mechanism that involves sequestering free heme into Hz. Investigations into the mechanism of formation of Hz are of major relevance to the pharmaceutical industry, which regards the inhibition of Hz formation as an ideal target for antimalarial drugs.84 Coronado et al.88 summarize the three main hypotheses for Hz formation: autocatalysis,89 biocrystallization, or catalysis by enzymes90 or lipids.91 Raman spectroscopy has played an important role in elucidating the mechanism of Hz formation because it provides information about the Hz crystals at submicrometer spatial resolution. For example, Egan et al.92 investigated the mechanism of the formation of Hz using different spectroscopic approaches, including FTIR and Raman spectroscopy. They demonstrated that the biomineralization of the β-hematin pigment takes place without catalysis by any enzyme and can occur near a lipid/water interface. Raman microspectroscopy was employed to monitor the formation of the pigment at the lipid/water interface. The results were supported with molecular dynamics simulations, which showed that in the absence of any competing hydrogen bond formation from the aqueous environment, it was possible to synthesize Hz. This confirmed the important role lipids play in the Hz synthesis inside infected RBCs. Nakatani et al.93 recently reported that histidine residues from the heme detoxification protein interact with the hemes and facilitate heme dimerization and consequently the formation of Hz. The above studies demonstrate how Raman spectroscopy can be used to investigate the structure and behavior of isolated Plasmodium metabolites. The approach can also be used to directly investigate cells in aqueous media due to the weak Raman scattering from water. For example, a typical Hz spectrum was obtained by directly targeting Hz crystals in the food vacuole of living P. falciparum trophozoites using a Raman microspectrometer.43,94 It was possible to differentiate the Hz

Figure 8. Average Raman spectra obtained with the use of 514.5 nm laser excitation from around 30 single functional, control RBCs (black) and 30 single iRBCs (red) measured in saline solution as described46 with the most prominent Raman wavenumber values (cm−1) for the overtones and combination modes for oxyHb (RBCs) and Hz observed inside iRBCs. The most prominent marker bands of Hz are marked with stars.

which are shifted with respect to predictions from their fundamental modes (see Table 1). The nonfundamental region of Hz does not obtain the strong band at 2890 cm−1 observed for oxyHb due to the fact that the ν11 mode is invisible in the fundamental region of Hz, so any of its overtone and combination modes of ν11 are also not present. 3.5. Mechanism of Hz Formation Studied with Raman Spectroscopy

Hz formation mechanisms have been widely studied, and nowadays, there are theoretical grounds for explaining the detoxification of heme by Plasmodium species, which is summarized in Figure 9.84 During the erythrocytic phase of the parasite’s life cycle, Hb is catabolized into amino acids, 5339


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Figure 9. Proposed representation of Hz formation within the intraerythrocytic life cycle of P. falciparum. Host Hb is taken up by the parasite and transported to the digestive vacuole through the cytostome. In this acidic organelle, the Hb is digested into small peptides and four toxic heme units (ferriprotoporphyrin IX). Neutral lipid bodies mediate the detoxification of the heme byproduct through the formation of Hz. Reproduced with permission from ref 84. Copyright 2013 Newlands Press Ltd.

Figure 10. SEM images of Hz and different types of Hz-like crystals. Synthetic Hz (a), natural Hz purified from P. falciparum culture (3D7) (b), and Hz-like crystals (c); FTIR spectra of hemin, β-hematin and HLC (d). Vertical arrows indicate typical bands associated with β-hematin and nHz, which are practically absent in the spectrum of HLC. Raman spectra of hemin, β-hematin, nHz, and HLC (e). Spectra were obtained using 632.8 nm laser excitation. The spectral lines of β-hematin and nHz are identical, while HLC shows a different pattern. Reprinted with permission from ref 99. Copyright 2015 BioMed Central (CC BY 4.0).

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Table 2. Studies on the Effects of Antimalarial Drugs Using Raman and Vibrational Techniques article

drug

technique

interaction

media

Frosch et al.107 Frosch and Popp105 Webster et al.111 Asghari-Khiavi et al.113 Alumasa114 Goto and Caseli115 Kozicki et al.112 Wood et al.51

chloroquine halofantrine chloroquine chloroquine, quinacrine, quinine and mefloquine quinine mefloquine chloroquine chloroquine

Raman (UV excitation) Raman (UV excitation) Raman Raman (NIR excitation) and IR IR PM-IRRAS ATR and Raman TERS

isolated isolated Hz ferriprotoporphyrin IX heme cell membranes Hz and lipids Hz

physiological conditions solid iRBC solid solid air−water interface iRBC iRBC

from the normal Hb within RBCs using Raman imaging combined with hierarchical cluster analysis.95 Ostera et al.96 recorded spectra of isolated food vacuoles and showed evidence of nitric oxide (NO) generated in situ, indicating that the NO form of ferrous heme nitrosyl complexes can influence the heme solubility inside the food vacuole. Hobro et al.97 studied the response of the macrophage after phagocytosis of the Hz pigment and demonstrated the difficulties encountered by the cells eliminating Hz crystals. The inhibition of the Hz synthesis is so important that an assay based on artificial Hz-like crystals (HLC) has been proposed for investigating the inhibition capabilities of drugs.98 The method is based on growing crystals of HLC, a compound very similar to Hz, under incubation at specific conditions. Studies performed by Tempera et al.99 using Raman and infrared spectroscopies showed that HLC is slightly different structurally to the natural Hz (nHz) extracted from parasite cultures and to synthetic β-hematin. Figures 10a−c show electron microscope images of the nHz, β-hematin, and HLC, respectively. nHz and β-hematin consist of needle-like crystals, the former being 700−1000 nm long and 60−90 nm wide and the latter 350−100 nm long and 150−250 nm wide with a squarer profile. In contrast, HLC crystals are organized around a central point in a 3−4 μm flower petal shape. Differences were also observed in the IR spectra (see Figures 10d and e), where HLC does not present the typical Hz carboxylate and C−O stretching bands at 1660 and 1220 cm−1, respectively. In the Raman spectra, there were similarities found between the HLC and the nHz in the ν15, ν10, and ν5 bands, but some of the other bands present in nHz were absent in the HLC. The results indicate there are distinct differences in the chemistry of HLC and Hz.

Figure 11. NanoRam hand-held Raman spectrometer. Reproduced with permission from ref 104. Copyright 2015 BioMed Central (CC BY 4.0).

their transparent blisters and obtained sensitivity and specificity values of 100 and 96%, respectively. In terms of drug investigations, the most studied interaction is the binding of chloroquine with Hz. Frosch et al.105−109 performed a number of studies investigating the structure of antimalarial drugs, including halafantine,105 the antiplasmodial naphthylisoquinoline alkaloid dioncophylline A,106,109 quinine in cinchona bark,107 and mefloquine.108 UV-based resonance Raman spectroscopy has been applied to investigate drug− protein systems, including dihydrofolate reductase with its inhibitor trimethoprim, gyrase with novobiocin, and catechol O-methyltransferase with dinitrocatechol.110 In these systems, the drug−protein interactions could be monitored by using excitation wavelengths in the vicinity of electronic absorption bands of aromatic amino acids and observing changes due to hydrogen bonding and/or possible dipole−dipole and dipole− polarizability interactions with the ligand. Webster et al.111 applied resonance Raman spectroscopy to detect structural changes in Hz following incubation with chloroquine. Although the chloroquine was not directly detected, changes in the relative intensity of Hz bands were observed after incubation with the drug for 24 h. In another study, Webster et al.82 recorded resonance Raman spectra using 514 nm excitation to demonstrate that chloroquine (CQ) acts as a molecular spacer and binds noncovalently through dispersion interactions with the hematin dimer, giving rise to π−π interactions between micro-oxo dimer units of Fe(III)PPIX, as evidenced by the decrease in intensity of ν4 in the Raman spectrum as a function

3.6. Raman Studies on the Structure and Effect of Antimalarial Drugs

Table 2 summarizes several studies investigating antimalarial drug interactions and structure with vibrational spectroscopy. Most of the drugs employed as antimalarials show strong Raman scattering, so this technique is ideal to (i) perform structural studies with and without the presence of Plasmodium sp. metabolites and (ii) detect and quantify the drugs under different conditions. For example, a number of studies have investigated the potential of Raman spectroscopy to detect counterfeit antimalarial drugs.100−103 Although an in-depth analysis of counterfeit drugs is out of the scope of this review, we highlight two interesting examples. Loethen et al.101 applied Raman spectroscopy to identify counterfeit antimalarial drugs using an in-house spectral library they developed. Furthermore, Visser et al.104 evaluated the potential of a hand-held NanoRam instrument (see Figure 11) for the analysis in situ of antimalarial samples. The authors evaluated 289 samples in 5341


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Figure 12. Confocal Raman imaging of malaria infected RBC. (a) A microphotograph of the measured infected RBC on the level of Raman stack measurement. (b) Autofluorescence of the RBC. (c) Integration maps of a bands assigned to the oxyHb or malaria pigment Hz in the region 1700− 1500 cm−1. (d) KMC results with the four main classes for chosen RBCs. Classes include Hz (yellow), oxyHb (red), low density heme (gray), and area around the Hz (pink). The black class observed around the RBC corresponds to a substrate signal and was removed as an outlier. (e) Average spectrum from each of the respective four classes of an infected RBC obtained from the KMC. Sampling densities were equal to 330 nm. The yellow color in panels b and c corresponds to the highest relative intensity of the compounds and was normalized for all cross sections to the most intensive obtained from the 0 μm optical section. In panels a−d, the imaged stacks cover the volume of 11× 11 × 9 μm3 (33 × 33 × 11 points).

and inside the digestive vacuole of a malaria parasite at 20 nm lateral resolution.49 However, no evidence of CQ directly binding to the Hz was found, which may be explained by either not having the required lateral resolution to detect the drug binding to the surface of the Hz crystal or sufficient signal-tonoise ratio for the measurement or, alternatively, the drug did not bind to the crystal face.

of increasing CQ mole fraction. In comparison, electronic spectra showed that CQ can bind to the unligated face of Fe(III)PPIX−OH/H2O monomers, potentially reducing the formation of π−π dimers.82 Kozicki et al.112 applied Raman and ATR-FTIR spectroscopic techniques to investigate malaria infected RBCs incubated with CQ. Cells treated with CQ showed higher levels of oxygenated hemoglobin (oxyHb) compared to untreated controls when using a combination of Raman microscopy and PCA. In summary, the majority of vibrational spectroscopic studies investigating drug interactions in malaria infected cells shows changes occurring within the cell metabolome. The direct detection of the drug in the cell is challenging because of its low concentration. The limitations in lateral resolution of conventional Raman microspectroscopy can be overcome by using near-field techniques. A first attempt116 by the Monash group combined the use of Raman and AFM to assess the effect of the different fixation procedures on RBCs at a subcellular level. In general, it was found that a mixture of formaldehyde (3%) and glutaraldehyde (0.1%) in buffer was sufficient to retain the structural integrity of cells with minimal autofluorescence.113 This protocol was also suitable for RBCs infected with P. falciparum parasites and preserved the characteristic knob-like structures on the infected RBC surface.116 More recent efforts have applied TERS to probe Hz crystals inside infected RBCs in search for drug binding sites.51 By sectioning infected RBCs in LR White embedding medium, it was possible to record TER spectra of Hz crystals protruding from the embedding medium

3.7. 3D Visualization of Infected Red Blood Cells

With the development of confocal microscopes for fast mapping, it is now possible to optically section cells. Figure 12 presents an example of a stacked measurement of cross sections recorded from a single infected RBC. Each cross section was obtained from an 11 × 11 μm2 area with the sampling density of 0.33 μm. The distance between cross sections is equal to 0.7 μm. As the lateral and depth resolution is approximately 0.36 and 0.8 μm, respectively, due to the chosen laser wavelength and optics, the imaging parameters enabled information to be obtained from the entire volume of the infected RBC. The integration time for a single spectrum was set to 0.2 s. Raman measurements and data analysis were performed using WITec software (WITec Plus). Figure 12c shows Raman maps based on the integration of bands in the 1700−1500 cm−1 region mainly from porphyrin modes in oxyHb and Hz. The yellow color corresponds to the highest relative intensity of the integrated area (the signal was normalized in the stack of images to the strongest band between 1590 and 1560 cm−1). This enabled the 3D 5342


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Figure 13. 3D chemical map based on integrated area of the amide I mode showing a red blood cell infected with a P. falciparum trophozoite calculated from a 3D reconstruction of the Raman images obtained from a red blood cell infected with P. falciparum: (a) Integrated absorbance obtained from the band at (1626 cm−1). (b) Cluster image showing two classes assigned to Hb (green) and Hz (red). (c and d) Average spectra of the red and green classes, respectively.

four-dimensional array (x, y, wnm, z), where wnm are the wavenumber values for each spectrum associated with the x, y coordinates of each image, and z is the image number (i.e., 1− 5). For each image, the Raman intensity of each spectrum was summed over the 1680−1620 cm−1 range, and this information was saved into a 3-dimensional array (x,y,z), which contains the summed intensity value for each 3-dimensional coordinate. Each point in this 3-dimensional array was interpolated by a factor of 4 by using the interp3 linear interpolation function in Matlab. Use of the jet color-map built into Matlab, which colors the image based on the summed intensity value, generated the final color. Finally, the image was sliced along the x, y, and z axes and displayed. The aspect ratio was set to [1.0, 1.0, 0.25] to stretch out the z-axis. 3D images can also be reconstructed using unsupervised hierarchical cluster analysis. Figure 13B shows another example of a P. falciparum trophozoite inside a RBC fixed in methanol (recorded at North Eastern University, Boston, United States). iRBCs were immersed in PBS and measured with 633 nm excitation (HeNe, Melles Griot) using a 60×/NA = 1.0 (Nikon) water immersion objective. The power at the sample was approximately 5 mW. Spectra were collected with 1 s illumination and a step size of 0.33 μm. For the 3D image reconstruction, the focus was manually moved by increments of 0.5 μm, and 7 scans were collected per cell. The image was created in SCIRun from the Raman 3D stack using 128 pixel interpolation to generate a smooth image.

visualization of the distribution of Hz and Hb in oxyHb as well as the parasitophorous vacuole where low Hz/heme signal was observed. The 3D autofluorescence image, which manifests as an increased background signal, is shown in Figure 12b. Cluster analysis was performed after cosmic-spike removal and background subtraction. The Raman data were analyzed with k-means clustering (KMC) using the Manhattan distance algorithm and were factored into four components that encompassed the entire cell volume. Figure 12e shows mean spectra of each class. The spectra are non-normalized to show low intensity areas from which additional information about cell density can be obtained. KMC discriminated between Hz (yellow class) and oxyHb (red class). The main marker band of Hz obtained with the use of 532 nm excitation is observed at around 1629 cm−1, which can be distinguished from the oxyHb spectrum which has the v10 mode at approximately 1640 cm−1. The class of low-density heme originates either from partially degraded oxyHb (gray class) or regions corresponding to the parasite vacuole. The pink class, which surrounds the Hz, is observed for the five stacks obtained from the top of cell but is not observed in the integration maps of Hz/heme due to weak signal and is therefore assigned to the less dense area of Hz. The KMC results are in agreement with those obtained by generating images based on the integration of characteristic bands. Vibrational spectroscopy provides an excellent means to visualize the spatial location of macromolecules throughout the entire cell volume.117 Figure 13a shows a 3D image based on the integrated area of the porphyrin stretching modes for a functional RBC infected with a P. falciparum trophozoite. The individual sections can be stitched together using a Matlab from Mathworks (Natik, United States) function developed inhouse. Each image from a total of five sections was placed in a 5343


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Figure 14. (a) Sketch of the setup for hyperspectral Raman imaging. It consists of three light paths: The green path represents the laser illumination system, consisting mainly of a microscope system of the lens L2 and the objective, a 30 m long optical fiber with a square shaped core and a 532 nm laser coupled in by the lens L1. The focal lengths are 1 cm for L1, 6 cm for L2, and 1.8 mm for the 100× objective. An additional HeNe targeting laser with 633 nm can be used by rotating a flip mirror. The cleanup filter blocks the fluorescence and Raman light generated in the illumination fiber. The path of the collected stray light is colored in red, consisting of a second microscope with an objective and tube lens. A dichroic mirror and an edge filter were used to block the Rayleigh light. The blue path represents the normal white light microscope and includes a Koehler illumination system. The microscopic image from the sample is imaged onto the camera by the objective, the tube lens, and the flip mirror. A lateral resolution of 1.25 μm was achieved using a 100× magnification objective. (b and c) Raman images of infected erythrocytes collected with the setup shown and application of the algorithm. Raman images and white light images are shown for four erythrocytes for samples from 6 (b) and 32 (c) h after infection. For each developmental stage, four RBCs are shown. All images are normalized in an intensity range from 0 to 1. Reproduced with permission from ref 121. Copyright 2015 Elsevier.

4. DIAGNOSIS OF MALARIA USING RAMAN METHODS

blood screening. The main disadvantage of this technique is the time taken to analyze an entire smear in search of an infected cell. By combining partial dark-field microscopy to identify microcrystals, it is possible to accelerate the analysis by selecting RBCs suspected of infection for evaluation using Raman microscopy. Other possible techniques such as confocal reflectance and quantitative phase microscopy (QPM) can also be combined with Raman spectroscopy providing a multimodal system, which improves the diagnostic capability.126 Raman spectroscopy can also be applied to monitor the progression of the disease in the body, as shown in a study performed in mice by Hobro et al.124 This study was extended to the analysis of plasma, which resulted in greater sensitivity (the reported limit of detection was below 0.2% of parasitemia) due to the lack of the Hb background. An alternative method is Raman acoustic levitation spectroscopy (RALS) for studying dynamics of oxidation−reduction cycles of Hb in red blood cells and for determining Hz in crude extracted trophozoite lysates.32 This proof of concept study demonstrated that Raman spectra can be obtained from negligible amounts of sample. The acoustic nodes generate highly concentrated droplets of trophozoite lysate, enhancing the probability of detecting infected cells even using a portable

4.1. Use of Raman Microscopes on the Detection of Hz in RBC

The strong scattering from Hz has been critical in a number of Raman spectroscopy-based approaches to malaria diagnostics both in cells and serum.71,73,97,111,118−124 However, the dependence on Hz detection is somewhat limited for studies on peripheral blood where the number of trophozoites and schizonts that contain abundant Hz is very low. In peripheral blood, early stage rings and gametocytes are more common, but these contain little Hz; hence, Hz is not an ideal marker for malaria infection. A number of different measurement modes have been proposed for the diagnosis of malaria using Raman spectroscopy, some of which are detailed in Figure 1 and explained above. Raman hyperspectral mapping was used to image a population of cells in a wide field of view to detect the presence of Hz. Using this approach, it was possible to identify individual RBCs infected with the parasite based on the strong porphyrin stretching band at 1569 cm−1.125 Small inclusions (less than 300 nm) of Hz could be detected in ring stage parasites indicating the suitability of the approach for peripheral 5344


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Figure 15. A picture of a Graphium weiskei butterfly (a) and a graphical representation of the nanoconical structures found on the butterfly wings (b). The yellow represents the gold coating. Reproduced with permission from ref 129. Copyright 2015 Royal Society of Chemistry.

Figure 16. Detection of Hz in blood using SERRS. Magnetic field-enriched SERRS and SERRS spectra of β-hematin supernatant obtained by centrifuging (β-hematin at concentration of 10−4 M) at an excitation power of 0.1 mW. Magnetic field-enriched RRS and RRS spectra of the same βhematin supernatant at an excitation power of 10 mW (a). Reproduced with permission from ref 130. Copyright 2012 Society of Photo-Optical Instrumentation Engineers (SPIE) (b and c) SERS peak intensity at 1623 cm−1 as a function of parasitemia level using methods 1 and 2, respectively. The data corresponding to normal blood samples, labeled as NB, are added manually to facilitate comparison. The asterisks indicate parasitemia levels at which the Raman peak intensity at 1623 cm−1 was significantly different to that in the normal blood sample in a t test (p < 0.05 for a, and p < 0.001 for b). A blood sample treated by method 1 and then mixed with nanoparticles synthesized separately (d). The blood sample after nanoparticle synthesis treated by method 2 (e). Reproduced with permission from ref 122. Copyright 2015 Springer Nature (CC BY 4.0).

optical fiber-based instrument. RBCs can also be selected and confined using light by means of optical tweezers. This approach was used by Dasgupta et al.123 for differentiating between optically trapped malaria infected and healthy RBCs according to their Raman spectra. Bilal et al.120 demonstrated that Raman spectra of plasma can be used to distinguish healthy, malaria infected, and dengue fever infected patients. Raman biomarkers for malaria have been identified from

clinical samples by Kozicki et al.127 In that work, Raman spectra of blood samples obtained from patients diagnosed with malaria and healthy controls were compared, and it was found that the ratio of the intensity of some Raman bands were different. For example, the I1130/I1075 was larger for the malaria infected samples, which the authors attributed to a low value of trans lipid conformations. On the other hand, the I2930/I2850 ratio was enhanced in the spectra of the infected samples which, 5345


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rings using the second method. It can be seen how in method 1 the Hz inside the rings is released around the sample, while in method 2, the Hz is still concentrated inside the ring. This is possibly the reason why in the second method the sensitivity of the technique was enhanced, obtaining a detection limit of 2.5 parasites/μL (see Figure 16b). This is compared with a sensitivity of 500 parasites/μL in the first method (see Figure 16c). However, the second method also showed less reproducibility and robustness than the addition of lysed RBCs to the already synthesized nanoparticles. A critical review of their work on the application of SERS to detect single parasites in the ring stage is available in ref 135. The aforementioned studies targeted Hz as both a SERRS active molecule and malaria biomarker. A different approach was used recently by Ngo et al.136 It identified a specific DNA sequence of P. falciparum using a sandwich hybridization of magnetic beads loaded with capture probes, target DNA sequences, and SERS nanorattles (nanoparticles which contain a spherical shell encapsulating a freely moving core particle in a solvent)137 loaded with reporter probes. The approach is illustrated in Figure 17a. The parasite DNA sequence targeted

according to the authors, was caused by modifications in structures of the membrane induced by the presence of the parasite. New instrumental developments such as a fiber arraybased hyperspectral Raman microscope121 (developed by the University of Jena, Germany) have enabled the acquisition of a hyperspectral image of a RBC with acquisition times in the range of seconds. The approach consists of wide-field imaging using an optical fiber bundle, which performs the transformation on a two-dimensional fiber array (8 × 8) into a line of 64 fibers focused at the slit of the spectrophotometer (Figure 14a). The in-house built microscope utilizes a tailor-made illumination system, which shapes the Gaussian-shaped profiled incoming light to a top-hat intensity distribution. In addition, the high throughput system was coupled with an algorithm which provides an image of the Hz pigment within the cell, as shown in Figures 14b and c. The method was able to detect a small deposit of Hz after 6 h of infection and also showed how the Hz deposits increase over 32 h (c). 4.2. Applications of SERS, SERRS, and SORS for Plasmodium Detection

One of the main disadvantages of Raman spectroscopy is the low sensitivity due to the low Raman scattering cross section of a molecule, which is typically around the order of 10−30 cm2 per molecule.34 Resonance Raman as described above can increase the signal by some three orders of magnitude, while metallic nanostructures used in SERS can overcome the low sensitivity by enhancing the signal by many orders of magnitude. For heme pigments, the combination of the two approaches is known as SERRS, which provides optimum enhancement of spectral features.128 Three metal nanostructures used for enhancing the Hz signal are reviewed here: gold-coated butterfly wings, magnetic nanoparticles with iron oxide core, and nanorattles. The wings of the Graphium weiskei butterfly contain nanoconical structures which enable SERS enhancement upon applying a gold coating and can be used as a biological alternative to the traditional synthetic substrates (Figure 15). It was demonstrated129 that these structures, composed of gold coated chitin protein, can be used to enhance the signal of Hz and thus identify the presence of the Plasmodium sp. parasite. The technique was able to reveal typical Hz bands in early stage low concentration (0.005−0.00005%) parasitemia in lysed RBCs. A more orthodox approach has been proposed130 and optimized131 by Yuen and Liu using magnetic nanoparticles with a silver shell and an Fe3O4 core. The method takes advantage of the magnetic properties of Hz132 to further enhance the SERRS signal. The authors studied the interaction of β-hematin crystals and nanoparticles under magnetic fields. Figure 16 shows the SERRS signal after enrichment with the magnetic field was approximately five orders of magnitude higher than the regular RRS, while the enhancement in the absence of magnetic field was just three orders of magnitude. In a recent study,122,133,134 their group compared the capabilities of silver nanoparticles using two slightly different SERS methodologies. They studied the outcomes of the analysis by (i) creating the nanoparticles before mixing with the lysed RBCs and (ii) synthesizing the nanoparticles inside the Plasmodium sp. Figure 16d shows lysed RBCs and parasites mixed with the nanoparticles as a result of method 1. In contrast, Figure 16e shows intact parasites at the ring stage after the RBC were lysed with nanoparticles synthesized inside the

Figure 17. (a) The nanorattle-based DNA detection method using sandwich hybridization of magnetic beads that are loaded with capture probes, target sequence, and ultrabright SERS nanorattles that are loaded with reporter probes. A magnet is applied to concentrate the hybridization sandwiches at a detection spot for SERS measurements. (b) SEM image of nanorattles bound onto the surface of magnetic beads in the presence of complementary target sequences. (c) Almost no nanorattle found on magnetic beads in the absence of complementary target sequences. Reprinted with permission from ref 136. Copyright 2016 Elsevier.

binds to the ultrabright SERS nanorattle and to the magnetic bead, creating a sandwich that is preconcentrated in a detection spot using a magnetic field. Figure 17b shows the SERS nanoparticles attached to the magnetic beads under the presence of the target DNA as opposed to Figure 17a, where the DNA was not inoculated and cannot be found in any particle attached to the beads. The SERS signal is proportional to the concentration of specific DNA sequence, which connects 5346


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the SERS nanorattles to the magnetic nanoparticles in the measurement surface. After optimization of all the parameters involved, the technique achieved an impressive LOD of 100 attomoles for P. falciparum DNA. Raman spectroscopy offers the potential for in vivo analysis especially when using near-IR lasers or spatially offset Raman (SORS). In a SORS measurement, at least two Raman measurements are performed; one at the source and one at an offset position of typically a few millimeters away. The two spectra can be then be subtracted by a scaled subtraction process to produce two spectra representing the surface and subsurface spectra. The technique could potentially be used to penetrate through thin membranes such as the ear, eye, and lip to detect malaria. Indeed, there is a patent on using confocal Raman spectroscopy to detect malaria through the eye lens.138 Although not a vibrational technique, a microvascular microscope has been developed, with dual modes of operation that utilizes cross polarized reflectance and transmission illumination to detect optical birefringence and absorbance from Hz, respectively.139 The device was designed to image superficial vasculature in the human oral mucosa, which in the future could incorporate Raman spectroscopy and be translated directly to human subjects.

Table 3. Typical IR Bands Present in the Spectra of RBCs and Plasmodium Speciesa band (cm−1) 3600−3500 3490 3277 2956

amide A, ν (N−H) ν3, νasOH ν1, νsOH νasCH3

2922 2874

νasCH2 νsCH3

2852 1740 1715

νsCH2 νCO ester carbonyl B-DNA base pairing vibration (nCQO and nCQN) A-DNA base pairing vibration (νCQO and νCQN) νCO amide I ν2, δ (H2O) amide II δasCH3

1708 1660, 1710 1640−1660 1642 1545−1530 1457 1450 1400

5. FTIR SPECTROSCOPY AS A DIAGNOSTIC TOOL FOR MALARIA

1300−1250 1244 1240, 1225, 1215 1208 1080 900−1200

5.1. Assignment of FTIR Spectrum from Malaria Infected Cells

Figure 18 depicts the ATR-FTIR spectrum of packed human RBCs. The IR spectrum is dominated by the amide bands

major contributor

assignment

a

δ CH2 νCOO2 of fatty acids and amino acid side chains amide III νas PO2 A-DNA, B-DNA, Z-DNA, νasPO2 νC−O DNA, νsPO2 νsC−O, νsC−C

proteins141 water water lipids/ proteins142,143 lipids142,143 lipids/ proteins142,143 lipids142,143 lipids142,143 DNA144 DNA144 Hz140 proteins141 water proteins141 proteins/ lipids141 lipids143 lipids/ proteins143 proteins141 RNA144 DNA144 Hz140 DNA144

ν, stretching; δ, bending; s, symmetric; as, asymmetric.

eye. This necessitates the use of multivariate analytical procedures to isolate the signals of the parasitic components from the rest of the components in the RBC to develop predictive models. Pioneering work by Slater et al.140 revealed that Hz is comprised of porphyrin units linked between the central ferric ion of one heme to the carboxylate side-group of another. The IR spectrum of Hz has an intense absorbance band at 1664 cm−1 that is absent from the spectra of either hemin or hematin, which suggests the presence of a unidentate carboxylate coordination onto iron in this pigment. Another distinct IR absorbance band observed in Hz at 1211 cm−1 is assigned to an axial C−O stretching wavenumber value from the axial carboxylate group. As discussed in Section 3.4, Hz is a good marker for trophozoites but not a good marker for early stage rings and gametocytes normally found in peripheral blood. FTIR spectroscopy can be used to detect: (i) the presence of some unique compounds associated with the Plasmodium sp. such as DNA (not present in human erythrocytes) and Hz and (ii) the changes induced by the infection in the parasite-RBC system such as lipogenesis, where neutral lipid spheres are synthesized for glycolysis activation and Hz formation. Hence, the use of infrared is not limited to detecting the presence of Hz but also allows for the detection of all stages of the parasites life cycle. The contribution from these different macromolecules essentially generates a molecular phenotype that can be identified using multivariate methods.

Figure 18. ATR-FTIR spectrum of packed RBCs in MeOH obtained from a patient infected with P. falciparum. Note: Main bands have been labeled with a color according to their assignment: red for proteins, green for lipids, blue for carbohydrate, violet for DNA, and brown for Hz.

located at 3268, 1630, 1527, and 1302 cm−1 from proteins, mostly from Hb, which is the major component in RBCs. Membrane lipids also contribute to the spectra in the CHn stretching (3100−2750 cm−1) and CH2 deformation (1447 cm−1) regions. The C−O stretching vibrational mode appears as a broad set of bands in the 1200−900 cm−1 wavenumber interval and is proportional to carbohydrate concentration. The band at 1233 cm−1 is assigned to the phosphodiester stretching vibration associated with membrane phospholipids and DNA from the parasite. The bands are strongly overlapped, and unique molecules from the parasite such as DNA, Hz, or changes in the lipid profile (see Table 3) appear as shoulder features in the spectrum that are hard to observe with the naked 5347


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Figure 19. Detection of infected cells using a focal plane array infrared microscope: (a) visible image. (b−d) Integrated area of the whole, C−H, and amide A region, respectively. (e) Image of a RBC smear (integrated area of the amide A region) with the detected cells marked as red. (f) Average second derivative spectra of the selected infected and uninfected cells. Reproduced with permission from ref 61. Copyright 2016 Royal Society of Chemistry.

region (3800−2500 cm−1), the C−H stretching vibration region (2990−2850 cm−1), and the amide A area (3400−3200 cm−1), respectively. The IR image of the whole region clearly identifies the presence of the parasite as a “hole” inside the cell, which may be explained by a lack of Hb in the food vacuole. The lack of material in this region is also shown in the images when integrating the C−H and amide bands but is less apparent due to the baseline correction used to eliminate the scattering contribution. The detection of the parasite inside the cell by the IR FPA microscope is useful for research, but in practice it does not gain any advantage over visible microscopy using a Giesma stain. Figure 19e shows an IR image of an 84 × 84 μm area from a blood smear. The cells are well delineated and, by using a Matlab image processing routine for identifying circles, it was possible to automatically select individual cells for analysis. A Matlab routine for extracting the average spectra of the different RBCs was also developed. When extracted, the second derivative of the average spectra of 14 infected and 34 uninfected cells showed differences as displayed in Figure 19f. The bands with minima at 2870 and 2952 cm−1 (corresponding to maxima in the raw spectra) were stronger in the case of the infected RBCs, while the band at 3290 cm−1 showed higher absorbance for the normal RBC. This is explained by the differences between the protein to lipid ratio in the cells. A PLS-DA achieved an 87% classification rate, demonstrating the potential of the technique to identify and quantify infected RBCs in a blood smear. FPA images can be also combined with Raman spectroscopy to provide multimodal images containing an infrared and Raman spectrum for each pixel.147 This allows for the combination of the information provided by the IR and Raman modes. The main challenges in applying this combination are the differences in spatial resolution between the modalities, the complexity of extracting biological meaningful information from the large volume of data, and the

5.2. Synchrotron FTIR

Pioneer studies utilized the IR beamline at the Australian Synchrotron to investigate normal and malaria infected RBCs at all stages of the parasite’s life cycle.145 Single point measurements of infected cells at the schizont, trophozoite, and ring stages showed large differences in the C−H (3100−2800 cm−1) and CO (1750 cm−1) stretching regions compared to normal cells. Recently, the IRENI beamline (SRC, University of Wisconsin-Madison) was expanded to cover a focal plane array detector using a series of mirrored optics and used to analyze smears of infected RBCs.146 This enabled the acquisition of images of several cells in one image plane, and again, important bands were identified that could distinguish the different stages of the parasite’s life cycle. More importantly, the data from separate visits to IRENI using different batch cultures could be combined to generate distinct clustering of the different life cycle stages of the parasite using principal component analysis. 5.3. Focal Plane Array (FPA) Imaging Using a Conventional Source

The synchrotron source proved to be very useful in identifying spectral differences between infected and uninfected cells, but it is not suitable as a routine clinical diagnostic technique. A recent study61 demonstrates the potential of applying a conventional FTIR FPA microscope for detecting malaria infected RBCs in blood smears. The study was performed using inexpensive microscope glass slides employed for routine Giemsa staining in hospitals. Although the opacity of the glass hampers the acquisition of the spectra below 2500 cm−1, the information contained from just the C−H stretching bands and the amide A band enabled the discrimination of infected and noninfected cells. The results of this study are presented in Figure 19. The visible image of a cell infected with a P. falciparum parasite is shown in Figure 19a. False color images in Figures 19b−d show the intensity of the integrated absorption between the whole 5348


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Figure 20. Analysis of multimodal vibrational imaging of two RBCs, performed after creation of an extended data matrix containing both Raman and infrared spectra for each pixel. (a) Visible and cluster images of cell 1 (top) and 2 (bottom). (b) Average Raman spectra of each cluster for cell number 1 (top) and 2 (bottom). (c) Average infrared spectra of each cluster for cell number 1 (top) and 2 (bottom). Third PC score value (d) and loading (e) of the principal component analysis created from the 10 average spectra obtained from the UHCA of cell 1 and 2. Reproduced with permission from ref 147. Copyright 2017 Royal Society of Chemistry.

in a RBC has not been previously reported because of the lateral resolution limitations of IR. In this case, the Raman served as “flag” for locating the correct position of the Hz and enabled the acquisition of extra information by means of the FTIR spectrum.

difficulties in integrating the images. In a recent study, multimodal vibrational images were generated of RBCs infected with P. falciparum parasites in the trophozoite stage. The Raman sampling interval was selected specifically to acquire images with the same pixel size as the FTIR FPA, and the sample slides were glued to a holder to enable measurement of the same cells in the same orientation. Images were first analyzed individually and then registered on an extended matrix containing an IR and a Raman spectra for each pixel. KMC performed over the individual FTIR data was not able to locate the parasite position inside the RBC. Nevertheless, the location of Hz IR bands from malaria trophozoites was possible following the fusion of IR and Raman data. Figure 20 depicts the multimodal imaging analysis performed over two different cells, and the KMC images (see Figure 20a) identify the region where the parasite is located. The average Raman spectrum shows the distinctive signature of Hz with a band at 1595 cm−1 (see Figure 20b). In the case of the FTIR average spectra (see Figure 20c), several bands from the class corresponding to the parasite (red color) are different from the uninfected RBC (blue and black colors). PCA performed using the average spectra from both infected and uninfected cells enabled discrimination between the red and blue classes using the third PC (see Figure 20d). Loadings of this PC (see Figure 20e) indicated bands characteristic of the IR spectrum of Hz lead to the separation. The specific location of the IR Hz bands

5.4. ATR-FTIR

FTIR microscopes are not suitable for use as the basis of a malaria diagnostic in developing countries due to their high cost and a lack of the expertise required to operate them. In addition, IR microscope-based technologies are also clearly outperformed by Raman and enzymatic alternatives in terms of cost-effectiveness and miniaturization capability. By contrast, ATR-FTIR is a very simple and cost-effective way to obtain the infrared spectrum of a sample, which has great potential in providing phenotypic information from cells and tissues. Instead of analyzing cell by cell, the sample is deposited on the ATR crystal as a suspension, and the average spectrum of all the cells is collected in less than a minute (see Figure 21a). In a recent study, a multivariate regression model was used to determine parasitemia levels from IR spectra of RBCs fixed in methanol using partial least-squares regression (PLS-R).7 ATRFTIR spectra of RBCs infected at different stages showed significant differences, associated with their different phenotype (see Figure 21b). For diagnostic purposes, a series of samples with different concentrations of ring stages were studied. Most successful PLS-Rs were obtained when the whole range of 5349


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Figure 21. Diagnosis of malaria and parasitemia quantification using ATR-FTIR spectroscopy of RBCs fixed in MeOH: (a) Conceptual diagram of the study. (b) PCA scores plot along PC1 and PC2 of control (C), ring (R), trophozoite (T), and gametocyte (G)-infected RBC data sets. PCA was performed after calculating the second derivative in the region 3000−2800 cm−1. (c−e) Regression plots for calibration and validation sets for three ranges of early ring-stage parasitemia: (c) model 1 (%) 0, 10, 15, 20, and 30; (d) model 2 (%) 0, 1, 1.75, 2.5, 3, and 5; and (e) model 3 (%) 0, 0.00001, 0.005, 0.01, 0.05, 0.1, 0.2, 0.4, 0.5, 0.8, and 1. Reproduced with permission from ref 7. Copyright 2014 Royal Society of Chemistry.

features in the IR (see Figure 22a). Sodium citrate presents strong absorption at 1572 cm−1 due to the carboxylate groups. Similarly, EDTA carboxylate groups are located at 1618 cm−1. Heparin presents also several bands in the spectrum assigned to carbohydrates (1146 and 1036 cm−1), secondary amines (1438 and 1420 cm−1), and sulfonated amine (1628 cm−1). All these contributions can interfere with the biological bands and hamper the diagnosis of malaria. In that study, blood samples were collected in EDTA, heparin, and citrate coated tubes, and ATR spectra of these samples were collected and compared. Unsupervised analysis (see score plots of a PCA in Figure 22b) showed that samples were clustered according to the anticoagulant in the tube, indicating that the anticoagulant is an important source of variance in the spectrum. Samples spiked with different levels of parasitemia collected in the different anticoagulants were also measured. The performance of PLS-R models for predicting the amount of parasitemia using ATR-FTIR were strongly affected by the anticoagulant, and only heparin showed successful calibration (R2 = 0.92) in the range 0−0.1%. This was consistent with the observation that heparin contributed less to the spectrum and is present at lower levels in the tubes. Other important sources of variability in real samples are the changes that accompany the metabolic composition of the samples. This is especially important when analyzing blood instead of packed RBCs. The analysis of whole blood eliminates a step of the process, but the metabolites present in serum contribute to the IR spectrum and can interfere with the parasite signal. On the other hand, malaria sickness may induce metabolic changes in plasma149 which can be targeted as markers if they provoke enough variations in the IR spectrum. In a recent study,150 the ability of the ATR spectrum to predict simultaneously urea, glucose, and malaria parasitemia from

concentrations was divided into short ranges of 0−30, 0−5, and 0−0.5% of parasitemia (see Figures 21c−e). The results indicated an excellent prediction capability of the parasitemia level using a cross validation method with the limit of detection calculated at 0.00001%, which is comparable to the gold standard of PCR. The regression vector of the PLS-DA model indicated that the models performed best in the CH2 and CH3 stretching region, which is consistent with the earlier work performed using synchrotron FTIR. The astonishing performance of ATR-FTIR in predicting the parasitemia level of laboratory samples opens the door to the development of a point-of-care methodology capable of use in the field at almost zero cost analysis. The proposed methodology faces two main challenges for its translation to the real world. First, the study published in ref 7 was performed using a Tensor 70 spectrophotometer from Bruker equipped with a MCT detector cooled with liquid N2. This and the large weight and volume of the spectrophotometer made its use as a point-of-care tool impractical. Second, it is likely that, compared to the artificial samples in the well-controlled environment of the cultural lab, real samples will show much more interference due to biological variability of blood components and preprocessing. More recent studies have tried to overcome the lack of portability using small and portable spectrophotometers such as the Agilent 45 or the Spectrum 2 from PerkinElmer, which can be transported to remote villages and can be battery operated. Regarding the biological variability, there is the critical problem of the lack of real samples available in the countries where ATR-FTIR methodologies are studied. Nevertheless, further studies have aimed to extend this technique to real world samples. Martin et al.148 studied the impact of the different anticoagulants in the diagnosis of malaria using ATR-FTIR. Common anticoagulants show strong 5350


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Figure 22. Chemical structure and second derivative spectra of citrate, EDTA, and heparin anticoagulants extracted from blood vials. PCA scores for dry plasma spectra collected in blood collection vials containing SC, EDTA, or heparin. Reproduced with permission from ref 148. Copyright 2017 Royal Society of Chemistry.

6. CONCLUSIONS AND OUTLOOK

thick smears of untreated blood sample was tested. ATR-FTIR spectroscopy is well-known for its capability in predicting the concentration of major clinical parameters in whole blood, serum, and plasma,53,151 but the simultaneous prediction of an infectious disease and clinical parameters has not been previously tested. This is important considering that glucose and urea levels are affected in patients suffering from malaria. Infrared spectra of a few microliters of sample deposited onto a glass slide and allowed to dry were recorded of the upturned slides in direct pressurized contact with the ATR crystal (see Figure 23a). The deposition of the blood on glass facilitated the acquisition, measurement, and storage of the samples. Results indicated that 5 μL of isolated red blood cells was enough to cover the entire ATR crystal with sufficient thickness to avoid any glass contribution to the spectra. Figure 12b depicts all of the 127 spectra of the database, and only 2 of them showed the characteristic Si−O bands of glass. Blood samples from controls were spiked with randomly assigned concentration levels of parasitemia (0−5%), glucose (0−400 mg/dL), and urea (0− 250 mg/dL). The root mean square error of cross validation (RMSECV) for parasite concentration was 0.58%, and for glucose and urea the relative RMSECV was 16 and 17%, respectively.

The work reviewed herein shows how vibrational spectroscopy can play an important role in the global fight against malaria. Raman and IR spectroscopy detect changes based on the molecular phenotype of the RBCs due to the presence of the parasite. The interpretation of these changes enables us to (i) study the molecular changes that occur during the development of the parasite throughout the erythocytic cycle, (ii) monitor the effect that antimalarial drugs have on the malaria parasite and host RBC, and (iii) diagnose even subclinical levels of malaria infection. The main components of the parasite, including DNA, Hz, and lipids show distinct spectral features in the IR and Raman spectra that can be used for diagnosis and research. Hz shows an enhanced Raman signal due to resonance enhancement, while the FTIR spectrum of Hz also shows distinct bands, making Hz the main target in the range of drug studies. The main drawback of Hz is that it is only produced in significant amount in mature stages of the parasite, and not present significantly in peripheral blood, hampering its use as a biomarker for diagnosis using blood obtained from venepuncture. However, Hz bands are extensively used in the study of drug interactions with the parasite. Lipid bands are the major 5351


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underway. The ATR units along with a laptop and microfuge can be run using a 12 V car battery for up to 8 h without recharging in tropical remote villages. The aims of these studies are primarily to determine if the ATR technique has the sensitivity to detect asymptomatic carriers, which is critical in the elimination of malaria as these people act as a reservoir for the parasite. New developments in sample preparation and instrumentation will see further gains in sensitivity and specificity, and thus, the future is looking bright for spectroscopic-based point-of-care diagnosis of malaria infection.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. ORCID

David Perez-Guaita: 0000-0002-2640-2927 Katarzyna M. Marzec: 0000-0002-8098-4622 Corey Evans: 0000-0001-7243-5650 Jose Garcia-Bustos: 0000-0002-6310-796X Bayden R. Wood: 0000-0003-3581-447X

Figure 23. Simultaneous determination of parasitemia, glucose, and urea in thick smears dried on glass slides. (a) Photo of the glass slides containing the thick smear on top. The slide was placed upside-down on the ATR. (b) FTIR spectra of the sample data set showing three samples with an abnormal value of absorbance in the 1000−900 cm−1 region, caused by a bad contact that places the glass in the path of the evanescent infrared beam of the ATR. Reproduced from ref 150. Copyright 2017 American Chemical Society.

Notes

The authors declare no competing financial interest. Biographies David Perez-Guaita obtained his Ph.D. in Chemistry from the University of Valencia (Spain) in 2014 working in the “Solutions and Innovations in Analytical Chemistry” research group and immediately joined Monash Centre for Biospectrosocpy in Melbourne (Australia). He has worked in several international laboratories, including the “Laboratoire de Physicochimie de l’Atmosphère” in Strasbourg (France), the organic chemistry department of the University of Valencia (Spain), the “Institut de Chimie Moléculaire” of Reims (France), and the “Institute of Analytical and Bioanalytical Chemistry” in Ulm (Germany). His research focuses on the development of vibrational spectroscopy-based solutions to clinical problems, including point-of-care, nanoimaging, and chemometrics applications.

spectral markers used in diagnosis-based IR studies. Lipids synthesized by the parasites in peripheral blood stages (e.g., rings) differ in quantity and structure from those present in uninfected RBCs. These differences are present in the spectra in a subtle way, requiring multivariate statistical techniques to capture them. Another IR parasite marker in the peripheral blood is DNA, which shows distinctive phosphodiester bands, but due in part to its low concentration, it is also necessary to employ multivariate statistics to identify these signatures. Both Raman and infrared spectroscopy are versatile techniques that can be applied in different scenarios. In the field of antimalarial drug research, Raman microscopy is widely used to study changes in the phenotype of the parasites caused by drugs as well as the interaction of drugs with Hz. Portable ATR is more suited than the other spectroscopic techniques in a field setting and for detecting early stage malaria parasites, including ring and gametocyte stages. Raman spectroscopy has single cell resolution, but the traditional Raman equipment has limited use in the field because of the time taken to image a field of cells and the dependence on the detection of hemozoin, which is not present in significant amounts in early stage rings and gametocytes. Focal plane array FTIR imaging microscopy has the speed to analyze a field of cells in a clinically relevant time frame, but the instrumental expense is not appropriate for the developing world. The next phase in the development of vibrational spectroscopy in malaria diagnosis is to implement some of these methods into a field setting. To this end, a preclinical trial of ATR spectroscopy in Thailand using independent nonexpert operators to perform the measurements has begun, with the data transferred to Monash University for analysis. The results of this study, to be published, show how portable IR instrumentation can be employed for the rapid diagnosis at low levels of parasitemia in remote regions. Point-of-care trials in remote villages in Laos and the Papua New Guinea are also

Katarzyna M. Marzec received her Ph.D. in chemistry at the Faculty of Chemistry at Jagiellonian University, Poland under the supervision of Professor L.M. Proniewicz. In 2010 she joined the Jagiellonian Centre for Experimental Therapeutics (JCET) at Jagiellonian University and the group of Professor M. Baranska as a postgraduate researcher. Currently, she is an assistant professor and the Head of the Raman Spectroscopy Laboratory at JCET. She carried out part of her graduate research at University of Coimbra, Portugal, under the supervision of Professor R. Fausto and some of her postdoctoral work as a visiting scholar at Monash University, Australia (Go8 Fellowship) under the supervision of Professor B.R. Wood and at University of Illinois, United States (Kosciuszko Fellowship) under the supervision of Professor R. Bhargava. Her research concerns the application of vibrational spectroscopy supported by complementary techniques to tissues, cells, and biomolecules studies. Andrew Hudson (BSc Oxon, Ph.D. Toronto) is an Associate Professor in Biophysical Chemistry at the University of Leicester. He has a track record for bringing frontier technologies to bear on problems at the life−science interface. Using vibrational spectroscopy, he has provided mechanistic evidence for how small molecules (such as NO and CO) confer protection on cardiomyocytes via their interactions with heme proteins, has detected phase transitions in lipid bilayer structures on the same length scale as a single mammalian cell, and has reported on 5352


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scopy to diagnostic medicine. He typically coordinates experiments conducted by his biomedical partners with spectroscopic sample measurement and analysis done at the Monash University Centre for Biospectroscopy (where he is the codirector) and the Australian Synchrotron, acting as a linchpin in these operations. He is a Group Leader in the Department of Microbiology and the Monash Biomedical Discovery Institute where he directs research programs in hyperspectral imaging of the human lung and brain and biospectroscopy of human stem cells and is heavily involved in developing point-of-care diagnostics of infectious diseases using ATRFTIR spectroscopy.

the binary coalescence dynamics of liquid−aerosol microdroplets. Dr. Hudson’s research also includes work on optical manipulation, microfluidics, and single molecule fluorescence spectroscopy. Earlier in his career, he worked for SMEs in the biotechnology and optical telecommunications sectors. Corey Evans attended Monash University in Australia and completed his undergraduate studies in 1993 and then went on to obtain a Ph.D. in 1998. He was then a postdoctoral fellow at the University of British Columbia and at the University of Kentucky. He began his academic career at the University of Leicester in England in 2003 where he is currently an Associated Professor in Physical Chemistry. He has broad interests in rotational, vibrational, and electronic spectroscopic techniques, computational chemistry, and in their application in the understanding of atmospheric chemistry processes and in the finding of new species of astrochemical interest.

Jack S. Richards is an Infectious Diseases Physician at Royal Melbourne Hospital and Laboratory Head at Burnet Institute. He has a particular interest in vector-borne diseases and tropical and travel medicine. His research seeks to develop and validate novel diagnostic tests against infectious diseases, especially for use in low and middle income settings. He also studies the development of naturally acquired immunity to infectious diseases to facilitate the development of novel vaccine strategies. He has ongoing collaborations in Vietnam, TimorLeste, Papua New Guinea, and Sri Lanka.

Tatyana Chernenko has over eight years of experience in the fields from biotechnology to physical chemistry and pharmaceutical sciences, developing cutting-edge diagnostic tools and applications. Her attention is currently focused on development of flow cytometrybased diagnostic instrumentation, benefitting fields spanning from oncology to microbial research. Her previous work was focused on implementation of noninvasive spectroscopic imaging modalities on various biological systems as well as synthesizing biocompatible nanopharmaceuticals to follow their subcellular trafficking and drugload-release patterns.

Dean Andrew graduated from Queensland University of Technology in Brisbane with BSc (Hons) in 2008 and worked on a Chlamydia vaccine project for 5 years, exploring the potential of novel adjuvants and different immunization routes in a vaccine against Chlamydia infections in humans and koalas. Dean moved to the Burnet Institute in 2013 to continue work on vaccine adjuvants in the malaria field, investigating vaccine protection against the blood stage of the malaria infection. Dean is currently based at the University of Melbourne studying malaria gametocytes.

Christian Matthäus studied chemistry at the Technical University Berlin and the University of Oklahoma. He received his Ph.D. working in Professor Max Diem’s laboratories at the City University of New York. His Ph.D. work focused on infrared and Raman spectroscopy on eukaryotic cells and tissues. In 2006, the group moved to the Northeastern University in Boston, where he was offered a postdoctoral position. In July 2009, he returned to Germany and started working as a research assistant at the Leibniz Institute of Photonic Technology and the Friedrich Schiller University in Jena in the group of Professor Jürgen Popp. His main research interests are linear and nonlinear spectroscopic imaging techniques, mass spectrometry, and their application to biological samples.

David A. Anderson is Deputy Director (Partnerships) of the Burnet Institute, Melbourne, and Co-Head of the Burnet’s Global Health Diagnostics Laboratory. He has worked at the Burnet Institute as a full-time research scientist and leader since completing a Ph.D. in molecular virology in 1988, with a strong focus on translational research that has led to a number of diagnostic and vaccine products that are licensed or under commercial development. He is the author of more than 70 primary research papers and an inventor on 7 active patent families in the areas of diagnostics and vaccines.

Miloš Miljković received his Bachelor’s degree in chemistry from the Belgrade University in 1997. He joined the lab of Prof. Max Diem in 2001 and defended his Ph.D. thesis in 2006 at the City University of New York. In 2006, he moved to Boston and held positions of associate research scientist and research assistant professor at Northeastern University. From 2013 to 2015, Miloš worked as a consultant with numerous startups and universities. As of 2015, he has been employed by Tufts University as laboratory manager. Currently, his research focuses on the intersection of spectroscopy, atomic force microscopy, and nanophotonics.

Christian Doerig obtained his Ph.D. in molecular virology at the Swiss Institute for Experimental Cancer Research (ISREC) in Lausanne, Switzerland. During his postdoctoral training, he pioneered the study of protein phosphorylation in the human malaria parasite Plasmodium falciparum and is now a recognized leader in this field. He is “Directeur de Recherches” at the French biomedical research agency Inserm and Honorary Professor of the University of Glasgow, Scotland. He established the first Inserm unit in the UK, at the Wellcome Trust Centre for Molecular Parasitology, University of Glasgow, and subsequently in Switzerland at the Inserm-EPFL joint laboratory, Ecole Polytechnique Fédérale de Lausanne (EPFL). In 2011, he joined Monash University as Head of the Department of Microbiology, a position he held until 2015. He is now a Professor of Microbiology at the same department, where he pursues his research on the kinomics of malaria parasites. A recently emerged research topic in his laboratory is the signaling response of host cells to intracellular pathogens, which at the moment covers work in Plasmodium-infected erythrocytes as well as in Flavivirus and Wolbachia infections.

After 20 years of academic research at the City University of New York in vibrational optical activity (infrared vibrational circular dichroism and Raman optical activity), Max Diem started in a new field in 1996, namely medical vibrational spectroscopy. He was instrumental in developing sound methods of correlating spectral data and medicinal information, and moved to Northeastern University (Boston) in 2005 for further development in infrared microspectroscopy of cells and tissues and Raman spectral imaging at subcellular resolution. He developed and applilied unsupervised and supervised methods of image analysis and is the cofounder of a biotech company to commercialize spectral diagnostics. He has been a Professor Emeritus since 2017.

Jose Garcia-Bustos is an Associate Professor at the Parasitology Department of the Faculty of Veterinary and Agricultural Sciences of the University of Melbourne. He has also retained an affiliation with the Department of Microbiology of Monash University as an adjunct. Prior to becoming an academic researcher and teacher in Australia, he

Philip Heraud is an experimental biologist who combines skills in spectroscopy, biomedicine, and data analysis to translate biospectro5353


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Attenuated Total Reflectance Infrared Spectroscopy and Multivariate Analysis. Anal. Chem. 2014, 86, 4379−4386. (8) Harris, I.; Sharrock, W. W.; Bain, L. M.; Gray, K.-A.; Bobogare, A.; Boaz, L.; Lilley, K.; Krause, D.; Vallely, A.; Johnson, M.-L.; et al. A Large Proportion of Asymptomatic Plasmodium Infections with Low and Sub-Microscopic Parasite Densities in the Low Transmission Setting of Temotu Province, Solomon Islands: Challenges for Malaria Diagnostics in an Elimination Setting. Malar. J. 2010, 9, 254. (9) Sturrock, H. J. W.; Hsiang, M. S.; Cohen, J. M.; Smith, D. L.; Greenhouse, B.; Bousema, T.; Gosling, R. D. Targeting Asymptomatic Malaria Infections: Active Surveillance in Control and Elimination. PLoS Med. 2013, 10, e1001467. (10) Moody, A. Rapid Diagnostic Tests for Malaria Parasites. Clin. Microbiol. Rev. 2002, 15, 66−78. (11) Barber, B. E.; William, T.; Grigg, M. J.; Piera, K.; Yeo, T. W.; Anstey, N. M. Evaluation of the Sensitivity of a PLDH-Based and an Aldolase-Based Rapid Diagnostic Test for Diagnosis of Uncomplicated and Severe Malaria Caused by PCR-Confirmed Plasmodium Knowlesi, Plasmodium Falciparum, and Plasmodium Vivax. J. Clin. Microbiol. 2013, 51, 1118−1123. (12) Waltmann, A.; Darcy, A. W.; Harris, I.; Koepfli, C.; Lodo, J.; Vahi, V.; Piziki, D.; Shanks, G. D.; Barry, A. E.; Whittaker, M.; et al. High Rates of Asymptomatic, Sub-Microscopic Plasmodium Vivax Infection and Disappearing Plasmodium Falciparum Malaria in an Area of Low Transmission in Solomon Islands. PLoS Neglected Trop. Dis. 2015, 9, e0003758. (13) Bunaciu, A. A.; Hoang, V. D.; Aboul-Enein, H. Y. Vibrational Micro-Spectroscopy of Human Tissues Analysis: Review. Crit. Rev. Anal. Chem. 2017, 47, 194−203. (14) Clemens, G.; Hands, J. R.; Dorling, K. M.; Baker, M. J. Vibrational Spectroscopic Methods for Cytology and Cellular Research. Analyst 2014, 139, 4411−4444. (15) Matthew, J.; Baker, J. T. Using Fourier Transform IR Spectroscopy to Analyze Biological Materials. Nat. Protoc. 2014, 9, 1771−1791. (16) Ferraro, J. R.; Nakamoto, K.; Brown, C. W. Chapter 1 - Basic Theory. In Introductory Raman Spectroscopy, 2nd ed.; Academic Press: San Diego, 2003; pp 1−94. (17) McCreery, R. L. Introduction and Scope. In Raman Spectroscopy for Chemical Analysis; John Wiley & Sons Inc.: Hoboken, NJ, 2000; pp 1−14. (18) Butler, H. J.; Ashton, L.; Bird, B.; Cinque, G.; Curtis, K.; Dorney, J.; Esmonde-White, K.; Fullwood, N. J.; Gardner, B.; MartinHirsch, P. L.; et al. Using Raman Spectroscopy to Characterize Biological Materials. Nat. Protoc. 2016, 11, 664−687. (19) Lasch, P.; Naumann, D. Spatial Resolution in Infrared Microspectroscopic Imaging of Tissues. Biochim. Biophys. Acta, Biomembr. 2006, 1758, 814−829. (20) Krishnan, R. S.; Shankar, R. K. Raman Effect: History of the Discovery. J. Raman Spectrosc. 1981, 10, 1−8. (21) Müller, U. M. Diem: Introduction to Modern Vibrational Spectroscopy, J. Wiley, New York, Chichester, 1993, ISBN 0−471− 59584−5, 285 Seiten, Preis: £ 49.50. Berichte Bunsenges. Für Phys. Chem. 1994, 98, 1347−1348. (22) McCreery, R. L. Instrumentation Overview and Spectrometer Performance. In Raman Spectroscopy for Chemical Analysis; John Wiley & Sons, Inc.: Hoboken, NJ, 2000; pp 73−94. (23) Modern Techniques in Raman Spectroscopy; Laserna, J. J., Ed.; Wiley: Chichester, New York, 1996. (24) Nafie, L. A. Recent Advances in Linear and Non-Linear Raman Spectroscopy. Part X. J. Raman Spectrosc. 2016, 47, 1548−1565. (25) Verma, P. Tip-Enhanced Raman Spectroscopy: Technique and Recent Advances. Chem. Rev. 2017, 117, 6447−6466. (26) Zhang, X.; Tan, Q.-H.; Wu, J.-B.; Shi, W.; Tan, P.-H. Review on the Raman Spectroscopy of Different Types of Layered Materials. Nanoscale 2016, 8, 6435−6450. (27) Ember, K. J. I.; Hoeve, M. A.; McAughtrie, S. L.; Bergholt, M. S.; Dwyer, B. J.; Stevens, M. M.; Faulds, K.; Forbes, S. J.; Campbell, C. J.

worked for GlaxoSmithKline in drug discovery for neglected diseases in Tres Cantos, Spain, coming from EMBO and Fulbright postdoctoral fellowships at the Biozentrum of Basel University and The Rockefeller University in New York, respectively. He holds a Ph.D. in Biochemistry from the Universidad Autonoma de Madrid and a B.Sc. in Fundamental Biology from the Universidad Complutense, also in Madrid. Don McNaughton is an emeritus professor of molecular sciences. He completed a Ph.D. in microwave spectroscopy studying transient species of astrochemically important species at Monash University in 1981 and followed that with an SERC postdoctoral fellowship at Sussex University, UK in transient species and astrochemistry. He was appointed a lecturer at Monash University in 1988 and has pursued interests in high resolution infrared spectroscopy and more recently in the area of biospectroscopy. His current research is highly multidisciplinary and involves developing Raman and infrared micro- and nanospectroscopy and imaging techniques to understand and follow biological processes and disease at a molecular level in cells and tissue. He also runs a program aimed at generating and studying, by high resolution spectroscopy techniques, species of interest in understanding interstellar and atmospheric chemistry. Bayden R. Wood is the Director of the Centre for Biospectroscopy, School of Chemistry, Monash University, Australia. He has over 130 peer reviewed publications and 10 book chapters, H index = 40. He is a founding board member of the Clinical Infrared and Raman Spectroscopy (CLIRSPEC) network and is an editor for Nature Scientific Reports, Applied Spectroscopy, and Sensors. He was recipient of the 2014 Doreen Clarke medal for Applied Chemistry from the Royal Australian Chemical Institute. He is also an Alexander Humboldt Fellow and is a Fellow of the Royal Society of Chemistry. His vision is to develop a world leading laboratory specialized in translating spectroscopy to find solutions for the most devastating diseases on the planet, including malaria, HIV, HBV, HCV, and sepsis.

ACKNOWLEDGMENTS Funding was provided by the Australian Research Council (Future Fellowship FT120100926 to B.R.W.). K.M.M. acknowledges funding by the Polish National Science Center (UMO-2016/23/B/ST4/00795). We acknowledge Mr. Finlay Shanks for instrumental support. REFERENCES (1) World Malaria Report: 2016; World Health Organization: Geneva; 2016. (2) Phillips, M. A.; Burrows, J. N.; Manyando, C.; van Huijsduijnen, R. H.; van Voorhis, W. C.; Wells, T. N. C. Malaria. Nat. Rev. Dis. Primer 2017, 3, 17050. (3) Cowman, A. F.; Healer, J.; Marapana, D.; Marsh, K. Malaria: Biology and Disease. Cell 2016, 167, 610−624. (4) Kasetsirikul, S.; Buranapong, J.; Srituravanich, W.; Kaewthamasorn, M.; Pimpin, A. The Development of Malaria Diagnostic Techniques: A Review of the Approaches with Focus on Dielectrophoretic and Magnetophoretic Methods. Malar. J. 2016, 15, 358. (5) Wassmer, S. C.; Grau, G. E. R. Severe Malaria: What’s New on the Pathogenesis Front? Int. J. Parasitol. 2017, 47, 145−152. (6) Loy, D. E.; Liu, W.; Li, Y.; Learn, G. H.; Plenderleith, L. J.; Sundararaman, S. A.; Sharp, P. M.; Hahn, B. H. Out of Africa: Origins and Evolution of the Human Malaria Parasites Plasmodium Falciparum and Plasmodium Vivax. Int. J. Parasitol. 2017, 47, 87−97. (7) Khoshmanesh, A.; Dixon, M. W. A.; Kenny, S.; Tilley, L.; McNaughton, D.; Wood, B. R. Detection and Quantification of EarlyStage Malaria Parasites in Laboratory Infected Erythrocytes by 5354


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