Volume 47, Issue 4 p. 438-445
Curriculum Development
Free Access

Bacterial cellobiose metabolism: An inquiry-driven, comprehensive undergraduate laboratory teaching approach to promote investigative learning

Gopal R. Periyannan

Corresponding Author

Gopal R. Periyannan

Department of Chemistry and Biochemistry, Eastern Illinois University, Charleston, Illinois, 61920

To whom correspondence should be addressed. Tel.: 217 581 6379; Fax: 217 581 6764. E-mail: [email protected]

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First published: 28 March 2019
Citations: 4

Abstract

Technique-centered biochemistry or molecular biology undergraduate laboratory curricula do not offer significant opportunities for thoughtful, in-depth exploration of the science to foster investigative learning. To demonstrate inclusion of inquiry-driven laboratory experiments into the undergraduate biochemistry and molecular biology curricula, a comprehensive set of laboratory experiments, covering several principles of biochemistry and molecular biology, have been developed under a single theme. The laboratory curriculum described here comprehensively investigates bacterial cellobiose metabolism using multiple biochemical, molecular biological (RNA isolation, RT-PCR, PCR, and enzyme assay), and analytical techniques (High Performance Liquid Chromatography, NMR, spectrophotometry, and thin-layer chromatography) to explore the principles of metabolomics and genomics in a single undergraduate laboratory course setting using Caulobacter crescentus as the model organism. This laboratory module serves as a model for educators to develop easy-to-implement laboratory curricula incorporating contemporary biochemistry and molecular biology concepts and techniques to provide a course-based undergraduate research experiences (CUREs) with defined learning objectives. © 2019 International Union of Biochemistry and Molecular Biology, 47(4):438–445, 2019.

Introduction

Development of inquiry-driven laboratory experiments that are effective and easy-to-implement is necessary for the transformation of undergraduate biochemistry and molecular biology curricula to hone the scientific process skills of students 1, 2. In this regard, significant progress has been made in implementing inquiry-driven laboratory teaching in different sub-disciplines of life sciences recently 3-7. Undergraduate laboratory lessons that solely aim to teach specific biochemical techniques, such as gel electrophoresis or enzyme assay, would limit the learning outcome merely to the practice of the technique. Alternatively, techniques that are used in experiments for answering a broader scientific question would encompass the rapidly developing biomolecular concepts and technologies in the field. Furthermore, technique-centered laboratory education does not provide a conducive educational environment for more thoughtful and in-depth exploration of the science to promote investigative learning 8. Development of laboratory teaching contents that incorporate experimental techniques as methods of answering a broader biochemical or molecular biological question, rather than technical training, would provide opportunities for students to perceive the technique in a broader scientific context and train them to critically evaluate the merits and limitations of the chosen technique.

The set of experiments outlined here investigate the aspects of bacterial carbohydrate metabolism using multiple biochemical, molecular biological and analytical techniques to explore the underlying biochemical, metabolomic, and genomic principles within a single undergraduate laboratory course. The interconnected metabolic processes of bacterial carbohydrate utilization are investigated using Caulobacter crescentus as the model organism and cellobiose as the carbon source 9. The theoretical scientific background behind these experiments are the contents covered by any biochemistry, molecular biology, and analytical baccalaureate curricula, and the techniques used are generally employed in a research setting; thus, this exercise provides a comprehensive course-based undergraduate research experience (CUREs).

Incorporating model organisms in undergraduate laboratory curriculum provides abundant opportunities for educators to develop inquiry-driven laboratory lessons with broader scope. Identification of safe, easy-to handle, and cost-effective model organisms that have been well researched and rich in documented information is the key for the successful development of such curriculum. For example, C. crescentus meets all these criteria to be an excellent model organism for education. It is a nonpathogenic, oligotrophic, gram-negative bacterium found in soil, fresh water lakes, streams and marine environments 10, 11. Genome sequence analyses and published works of C. crescentus show genes for plant polysaccharides metabolism through a variety of special pathways, such as Entner-Doudoroff type glycolysis and pentose phosphate pathway and nutrient uptake 9, 12, 13. Furthermore, published data show that these highly regulated carbohydrate metabolic pathways strategically connect to environmental polysaccharide degradation mechanisms to use the scarcely available polysaccharide fragments as carbon sources in an oligotrophic environment 12.

These genetic, metabolic, and nutrient acquisition concepts can be incorporated into related lecture contents as special topics for discussion in advanced biochemistry or molecular biology courses. Availability of biochemical and molecular biological experimental tools (vectors, mutants, etc.) and well-annotated genomic, proteomic, and metabolomics databases related to C. crescentus makes it an easily adoptable, safe bacterial model organism with broader educational outcomes even with limited resources 13. Here, it is demonstrated how comprehensive interactomic data can be gathered on the networks of enzymes and metabolites for cellobiose metabolism by C. crescentus, and how discussions can incorporate the roles of transporters and regulators to provide a broader content exploration opportunity in the areas of biochemistry, molecular biology, genomics, and related bioanalytical techniques.

The recent revision of the Medical College Admission Test (MCAT) by The Association of American Medical Colleges includes several changes to the required core competency for premedicine students with significantly broadened biochemistry content, underscoring the necessity of comprehensive biochemical knowledge for medical students. As per these changes, The Biological and Biochemical Foundations of Living Systems section of the MCAT requires students to demonstrate skills that reflect their knowledge in scientific inquiry and reasoning, research methods, and statistics. Furthermore, the American Society for Biochemistry and Molecular Biology, which was tasked with the development of the MCAT changes, recommends biochemistry laboratory educators to include experiments to develop these skills through hands-on experience 8, 14. Introduction of laboratory courses that incorporate the elements of biochemistry-related scientific inquiry would benefit the premedicine students as well as others. However, development of such laboratory content remains challenging due to limited institutional resources and logistical issues of implementation. The primary aim of this tested laboratory curriculum is to serve as a model to provide biochemistry and molecular biology educators with a suitable inquiry-driven laboratory exercise with a broader scope. This inquiry-driven curriculum can be taught at institutions with limited facilities by modifying the experimental goals that fit to the available resources.

Conceptualization of this inquiry-driven laboratory exercise originated from a publication from our research laboratory. We reported the gluco-oligosaccharide utilization by C. crescentus and found evidence for extracellular breakdown of gluco-oligosaccharides and cellobiose, as well as cellobiose uptake as a disaccharide across the membrane 15. This article by Presley et al. would serve as the source for materials and methods as well as a base for any modification of this content for those who may adopt this material for teaching purposes. This laboratory exercise was taught twice in successive years for 15 students in total. All experiments were designed and taught by the author as the instructor of the course with the help of a teaching assistant and a stockroom personnel in material preparation. Successful implementation of this laboratory exercise demonstrates the possibilities of transforming suitable published research projects into educational products with excellent learning outcomes.

Learning Objectives

The target student groups are junior and/or senior (i.e., third- and fourth-year students in a four-year program) undergraduate biochemistry majors or students who have completed a typical biochemistry lecture series or a combination of biochemistry and molecular biology courses. These students are expected to have significant, if not all, theoretical background knowledge on the biochemical, molecular biology, and analytical principles of these laboratory exercises. The goals of this laboratory design are the following: first, to promote student-led, inquiry-driven planning and performance of experiments in a collaborative manner to address a broader research question. Second, to provide the necessary technical and hands-on experience on many biochemical and molecular biological techniques that is typically taught in a traditional biochemistry laboratory course. The following specific learning objectives guide the students to develop their experimental design ability upon completion of this laboratory exercise:
  1. Comprehend a research question and then plan and choose appropriate experiments with suitable technique/s to answer the question.
  2. Connect and apply theoretical biochemical and molecular biological knowledge to a practical problem.
  3. Demonstrate proficiency in standard biochemistry and molecular biology laboratory techniques to collect and analyze data and critically evaluate the efficacy of the experimental design with respect to the scientific literature and make necessary changes to the scientific process.
  4. Demonstrate ability to communicate and collaborate in a productive manner to achieve intended goals.

Infusion of these essential elements of scientific research practices into undergraduate curriculum would elevate the level of training and quality of education necessary to prepare competent future researchers, educators, and health professionals to succeed in a competitive environment.

Laboratory Format

This laboratory exercise was presented to the students in the form of research questions, deviating from the traditional biochemistry laboratory teaching methods, to test the following general hypothesis: Caulobacter crescentus can utilize cellobiose as carbon source for growth by activating the transcription of glycoside hydrolases to break down cellobiose extracellularly. Students worked in pairs with the permission to freely share and discuss the outcome of each other's experimental results to learn how research teams would work collaboratively to achieve the intended research goals. There were four groups of two students and each student group conducted all the experiments to comprehensively test the hypothesis within the group. A typical laboratory period included a prelaboratory assignment (assigned as homework during the previous laboratory period), prelaboratory discussion, laboratory procedure and a postlaboratory quiz.

To promote independent thinking and scientific creativity, each of the research questions was presented as a prelaboratory assignment to comprehend the research question and develop an outline of experimental method/s to find answer/s. During the prelaboratory, the whole class actively discussed their thoughts and plans as a single group with the facilitation of the instructor. Following the discussion, with the appreciation of students' suggestions, students were given the procedure/s to conduct the experiment/s. For some questions, the experimental procedure was exactly what students proposed but for other questions, it was different as the experimental design was dependent on the laboratory resources available. Upon completion of a laboratory procedure, students individually answered a prelaboratory quiz to evaluate their own comprehensive understanding of the question and experimental design in a broader biochemical and molecular biological context.

Experimental techniques employed in this exercise included several biochemical, molecular biological, and bioanalytical techniques that are typically taught in a biochemistry/molecular biology laboratory course as well as few advanced techniques based on instrument availability. The biochemical and molecular biological techniques included: RNA isolation, reverse-transcription, polymerase chain reaction, gel electrophoresis, cellular fractionation and enzyme isolation, and enzyme activity assay. The analytical techniques included: 1H-NMR, High Performance Liquid Chromatography (HPLC), thin-layer chromatography (TLC), and UV–Vis spectroscopy. Advanced techniques that may be available at research-intensive institutions, such as proteomics, RNA-Seq or RNA microarray and mass spectrometry, can also be included to introduce cutting-edge technologies with a broadened scope of this laboratory design. In addition to these biochemical and analytical techniques, bioinformatics analyses were used to aid in answering some of the questions and to emphasize the efficient use of bioinformatics in research.

Research Questions, Methods of Investigation, Representative Data, and Learning Outcomes

The description below summarizes the experimental methods and a representative data collected by students for each question. For each research question, the technical and critical thinking aspects of the learning objectives achieved are summarized. Primary literature citations and other resources are provided to aid in the reproduction of these experimental methods.

Question 1: Can Caulobacter crescentus utilize cellobiose as a sole carbon source for growth?

Methods Summary: Caulobacter crescentus (CB15 strain) cultures were raised in the presence of cellobiose as the sole carbon source in M2 minimal medium. Optical density (OD600) of the cultures was measured using a spectrophotometer to monitor cell growth as a function of growth time (Fig. 1). Detailed experimental conditions can be found in the Supporting Information section and elsewhere 13, 15.

Details are in the caption following the image
Comparison of C. crescentus growth measured by OD600 in M2 minimal medium supplemented with 0.2% (w/v) glucose or cellobiose. Cultures were grown at 30 °C in a 260 rpm incubator shaker.

Results

Learning Outcomes:

Technical skills developed: 1) Establishment of bacterial cell cultures; 2) measurement of optical density-based bacterial growth; 3) graphical presentation of the growth data as a function of time.

Critical thinking: 1) significance of the type of carbon source in bacterial growth rate: why would a monosaccharide and a disaccharide give two different growth curves? 2) Time line of metabolic transformation: why did growth with cellobiose show an extended lag phase (approximately 10 hours) compared to glucose? 3) What is the significance of minimal media nutrients for bacterial growth and propagation?

Question 2: Does Caulobacter crescentus extracellularly break down cellobiose or metabolically transform cellobiose, or uptake cellobiose across the membrane?

Method Summary: The extracellular medium was analyzed to identify the chemical nature of the transformation of cellobiose carbon source as a function of growth time using HPLC and 1H-NMR spectroscopy 16, 17 (Fig. 2). In place of these techniques, simple TLC can be used for the same effect as demonstrated below (Fig. 2) 18.

Details are in the caption following the image
(a) HPLC analysis of the cellobiose extracellular medium after 12 hr growth period. Glucose and Cellobiose were detected as ELSD response (b) 1H-NMR analysis of the extracellular medium collected over 48 hr growth period. (c) TLC analysis of the cell-free growth media collected after 12 hr of growth are shown with corresponding standards.

Results

Learning Outcomes:

Technical skills developed: 1) handling of bacterial cultures, centrifugation, lyophilization, preconcentration; 2) HPLC/1H-NMR/TLC: sample preparation, method development for analyses, and instrument use. Method development for each technique included the following specifics: a) TLC: determination of proper solvent, determination of standards, and orcinol-H2SO4-based carbohydrate detection method; b) HPLC: determination of HPLC column and suitable solvent for the separation of the analytes (glucose, cellobiose, or derivatives), development of an elution profile, optimization of sample concentration, column temperature and detector response ELSD or Refractive Index (RI) Detector; c) 1H-NMR: preparation of a solvent system and reference, optimization of spectral parameters; 3) comparative analyses and interpretation of data. For example, a) TLC: calculation of Rf values of standards and analytes, b) HPLC: chromatographic analysis in terms of retention time and peak area, c) 1H-NMR: spectral analysis in terms of chemical shifts and coupling constants.

Critical thinking: Upon completion of the data analyses, students answered questions related to metabolism in conjunction with the growth pattern observed in the previous experiment. Additionally, students critically evaluated the appropriateness and the pros and cons of the analytical techniques employed. Some discussion topics involved were 1) relationship between observed carbon compound and growth pattern: what chemical transformation did cellobiose undergo and at what time point in the growth did the metabolic transformation occur? 2) Collective analyses and interpretation of data to derive conclusions about cellobiose utilization, and 3) suitability of analytical techniques: were the techniques ideal for glucose/cellobiose analyses? What are the alternative techniques suitable for the analysis of cellobiose, glucose, and their derivatives?

Question 3: Which glycoside hydrolases may be responsible for the breakdown of cellobiose?

Method Summary: Exploration of carbohydrate metabolic pathways using metabolic databases of C. crescentus to identify and list all putative glycoside hydrolase (GH) involved in the breakdown of cellobiose (Fig. 3). The instructors and/or students can identify several databases but perhaps the most useful ones for this laboratory exercise that are publicly available would be KEGG Pathway (http://www.kegg.jp/kegg/kegg2.html) 19, the Carbohydrate-Active enZYmes database (www.cazy.org) 20, and the genomic information site from NCBI (https://www.ncbi.nlm.nih.gov/genome/?term=Caulobacter+crescentus) 9. Additionally, the bacterial subcellular protein localization prediction tool PSORTb (version 3.0.2) (https://www.psort.org/psortb/) can be used for the prediction of subcellular location of the identified GHs to generate a table as shown by Presley et al. 15, 21.

Details are in the caption following the image
Publicly available databases: (a) CAZy and (b) KEGG pathways were used to explore and identify the glycoside hydrolase enzymes involved in the breakdown and conversion of cellobiose in Caulobacter crescentus. [Color figure can be viewed at wileyonlinelibrary.com]

Results

Learning Outcomes:

Technical skills developed: 1) identification of databases that are suitable for the study and execution of searches with appropriate technical criteria; 2) verify accuracy of the information across multiple databases; 3) gather information that is relevant to the investigation, compile the data, and further process.

Critical thinking: 1) Why does an organism need to code for several GHs? 2) What are some possible differences or distinction among these GHs? 3) What is the significance of GH-specific subcellular location of GHs in the metabolic conversion pathway of cellobiose and how it is related to other enzymes and metabolites of cellobiose/glucose metabolism? (iv) How does the subcellular localization prediction fit with the data obtained in the preceding experiments?

Question 4: Which of the predicted glycoside hydrolase genes are expressed for the conversion of cellobiose?

Method Summary: To identify the GHs responsible for the breakdown of cellobiose into glucose by C. crescentus, the nine GH genes listed in the table given in the Supporting Information Materials were tested for expression using reverse transcriptase technology (Fig. 4). This part of the gene expression analysis involved three steps that can be treated as separate experiments: 1) RNA isolation using TRIzol® Max™ reagent 22, 23; 2) cDNA synthesis using reverse transcriptase enzyme; 3) PCR amplification of the cDNA products 15. Additional information is provided under Supporting Information Materials section.

Details are in the caption following the image
RT-PCR based glycoside hydrolase gene expression analysis of C. crescentus. Cells collected after 3 h growth in cellobiose were used for the isolation of total RNA and cDNA synthesis. C. crescentus glycoside hydrolase gene expressed in response to cellobiose carbon source was identified using gene specific primers and standard PCR techniques.

Results

Learning Outcomes:

Technical skills developed: 1) RNA isolation; 2) cDNA synthesis; 3) polymerase chain reaction; 4) agarose gel electrophoresis.

Critical thinking: 1) Why were only few GH gene expression detected; is it due to gene regulation? 2) Are all GH genes expressed in the same amount? What methods available for the quantification of gene expression.

Question 5: What are the primary subcellular locations of glycoside hydrolases that may be responsible for cellobiose breakdown?

The metabolic utilization of cellobiose by C. crescentus requires conversion of cellobiose into glucose monomers. This can be accomplished two ways: 1) extracellular breakdown of cellobiose into glucose or 2) by the uptake of cellobiose disaccharide across the outer membrane and further transformation in the periplasmic space and/or cytosol. Data gathered for Question #2 indicate that primary mode of cellobiose conversion is by extracellular breakdown into glucose.

Method Summary: Students used cellular fractionation protocols from published research articles 24, 25 to isolate extracts that may contain GH enzymes. Outer membrane and periplasmic fractions of C. crescentus grown in cellobiose was isolated and tested for GH activity of these subcellular fractions using p-nitrophenyl-β-glucoside (pNPG) as the substrate and UV–Vis spectroscopy (Fig. 5) 10. Total protein concentration of the subcellular fractions was determined using Bradford assay (Bio-Rad) as per manufacturer's instruction. Additional information are provided in Supporting Information Materials.

Details are in the caption following the image
Reaction scheme for the formation of pNP from pNPG catalyzed by GH.

Results

Total units and specific GH activities of periplasmic and outer membrane fractions were calculated using the change in absorbance and the equations given below (Table 1). These equations are essential components of enzymology-related experiments in any biochemistry laboratory education.
Activity = Amount of p NP produced Time 1 Unit of Activity U = 1 μmol = 1 μmol of p NP / min Relative Activity = Units U Enzyme Volume mL Specific Activity = U Amount of Protein
Table 1. Glycoside hydrolase (GH) activities of subcellular fractions of cellobiose culture
Enzyme source Δ Abs (@ 405 nm) Units of GH activity (U) Specific activity U/mg protein
Outer membrane 0.14 12.4 × 10−4 0.037
Periplasmic fraction 0.10 9.6 × 10−4 0.035

The above experiment of GH activity can be expanded to a substrate-specific cellobiose hydrolase activity assay by monitoring the generation of glucose by a coupled activity of glucose oxidase 26.

Learning Outcomes:

Technical skills developed: (i) Cellular fractionation; (ii) protein quantitation using Bradford assay that included generation of calibration curve; (iii) optimizing enzyme reaction and spectral conditions to make valid spectrometric measurements by choosing appropriate buffer, temperature, and suitable enzyme substrate concentrations and wavelength; (iii) UV–Vis spectrometry-based measurement of enzyme activity as a function of changing absorbance and calculation of enzyme activity.

Critical thinking: (i) How do different extraction conditions facilitate the selective fractionation of outer membrane and periplasmic contents in terms of structure and composition of these cellular compartments? (ii) How does the strategic localization of GHs in the periplasmic space help C. crescentus acquire cellobiose as a carbon source? (iii) How to develop an analytical method (in this case UV–Vis spectrophotometry) to measure a biochemical activity (in this case GH enzyme activity). What other alternative analytical techniques can be employed to measure GH enzyme activity?

Discussion

Undergraduate biochemistry and molecular biology laboratory curriculum is an ideal opportunity to expose students to the chemical basis of life to motivate them to probe the underpinnings of life as we know it. An investigative learning experience could be a suitable teaching approach to incorporate different curricular topics learnt in previous courses to motivate the students to pursue the fields of biochemistry or molecular biology as a career path. Novel investigative laboratory experiences require constant modification of the curriculum to include contents related to the rapidly progressing field of biochemistry and molecular biology. For this purpose, the biochemistry and molecular biology curricula need to be a fluid structure to adapt to the advances in the biotechnology fields that are easy to implement. This set of comprehensive experiments stands as an example to develop comprehensive investigative biochemistry and molecular biology teaching laboratory contents that can be implemented even at an undergraduate institution with limited resources.

The key to successful designing and implementing a comprehensive investigative undergraduate laboratory depends on the identification or formulation of a biochemical or molecular biology question that can be answered by a wide variety of experiments that include traditional and modern techniques. Published research work in biochemistry and molecular biology can be turned into teaching modules with modification to suit a teaching setting. For example, any research content that involves nonpathogenic bacteria or C. elegans as a model organism and explores fundamental biochemical or molecular biology phenomena could be transformed into effective laboratory teaching modules. Adoptability of a research-derived set of experiments requires meticulous consideration, including but not limited to: modifiable to achieve the intended curricular goals, suitability to the student group, availability of laboratory resources and supplies, and flexibility to scheduling. Issues with one or more of these considerations may preclude educators to promote investigative laboratory curriculum and resort to cookie-cutter experiments 1. One way of seeking resources to include cutting-edge experiments is to seek the support of a research facility at a nearby research intensive institution. In the past, we have approached University of Illinois facilities (Protein Science) to conduct parts of 2D-gel electrophoresis experiments and microscopic imaging (Carl R. Woese Institute for Genomic Biology: Microcopy and Imaging Facility) in undergraduate teaching laboratory, and typically these institutions are receptive to promote such education missions as encouraged by federal grants.

Students enrolled in this course were junior and senior undergraduate biochemistry students, and almost all had completed two or more biochemistry and/or molecular biology courses as well as a course on quantitative analysis. Each set of investigative experiments included prelaboratory and postlaboratory worksheets/quizzes to provide an opportunity to revisit the related theoretical background and to assess the achievement of learning objectives upon completion of the experiments. A majority of the students enrolled in the course were aiming to pursue graduate studies in biochemistry or health-related fields, and four of them were pursuing Doctoral degrees in Biochemistry or Master's degrees in Biochemistry and Bioengineering. The higher level of enthusiasm and motivation exhibited by students may be due to their ambitions to pursue higher studies that were galvanized by this investigative learning opportunity.

A survey conducted showed increase in the confidence levels closer to 90% in all aspects of the learning objectives, indicating the increase in student confidence level in achieving the goals set for this exercise. A few of the students suggested providing more time for prelaboratory discussions and meticulous planning of the time allocation to complete all the required experiments.

In summary, inquiry-driven biochemistry and molecular biology laboratory teaching methods allow students to develop scientific reasoning and critical thinking skills and efficient learning of the scientific techniques and the methods of scientific data reporting. Transformation of suitable biochemical and molecular biological research outcomes into teaching materials provide superior learning opportunities for students to experiment contemporary technologies than traditional technique-based lessons as assessed by Goodey et al. and evidenced by this exercise 2.

Acknowledgments

Bacterial strain and technical assistance related to Caulobacter crescentus handling were kindly provided by Drs. Aretha Fiebig and Sean Crosson of University of Chicago. The author thanks the students of CHM3455 course at the Department of Chemistry and Biochemistry of Eastern Illinois University for their participation and contribution in generating some of the data shown in this work. The author acknowledges the help of Dr. Thomas Canam of Department of Biological Sciences, Eastern Illinois University for his careful reading and editing of the manuscript. The author thanks Redden Fund program of Eastern Illinois University for financial support.